Download This thesis of Michael Andrew Thompson Freiberg was reviewed

The Pennsylvania State University
The Graduate School
College of Agricultural Sciences
EXPLORING THE IMPACT OF THE ANTIVIRAL DRUG RIBAVIRIN ON RNA
VIRUSES IN HONEY BEES
AND
THE PRESENCE OF RNA VIRUSES IN BEES IN BRAZIL
A Thesis in
Entomology
by
Michael Andrew Thompson Freiberg
 2012 Michael Andrew Thompson Freiberg
Submitted in Partial Fulfillment
of the Requirements
for the Degree of
Master of Science
May 2012
ii
This thesis of Michael Andrew Thompson Freiberg was reviewed and approved* by the
following:
Diana Cox-Foster
Professor of Entomology
Thesis Advisor
Ed Rajotte
Professor of Entomology
Christina Grozinger
Associate Professor of Entomology
Craig Cameron
Paul Berg Professor of Biochemistry and Molecular Biology
Eddie Holmes
Professor of Biology
Gary Felton
Professor of Entomology
Head of the Department of Entomology
*Signatures are on file in the Graduate School.
iii
Abstract
Honey bees, Apis mellifera L., are essential pollinators worldwide and have an
enormous economic impact. Unfortunately, however, honey bee health is declining.
Honey bee viruses represent a significant threat to honey bee health and are associated
with a variety of pathologies such as dead brood, deformed wings, and general colony
mortality. Several viruses have even been associated with the recent mass die-off of
honey bees reported in North America called Colony Collapse Disorder. Despite the
threat posed by honey bee viruses, there are no chemotherapeutic agents currently in use
to treat these viruses and so treatment remains limited to management techniques which
are largely ineffective. Beekeepers are thus often forced to destroy infected colonies in
order to prevent the spread of viral diseases. I explored the broad-spectrum antiviral drug
ribavirin for its potential as an antiviral treatment in honey bees. I present evidence that
ribavirin was active against Deformed wing virus (DWV), Black queen cell virus
(BQCV), and Sacbrood virus (SBV). I also explored the mutagenic effect of ribavirin on
viral genomes. Although further study is required to determine if ribavirin is safe and can
improve colony survival, this study serves as an important step in the development of an
antiviral treatment for viral infections in honey bees and may also provide insight into the
ecology and epidemiology of these viruses.
In addition to this study on ribavirin, I explored the prevalence of honey bee viruses
in Brazil, both in honey bees and native stingless bees (Apidae; Meliponini). In North
America, honey bee viruses have been found in native bees such as the bumble bees
(Apidae; Bombus) where they have been shown to cause disease. Transmission can occur
through pollen between honey bees and native bees if both species are foraging on the
same flowers. The potential for this cross-species transmission poses a health risk to
native bees which are likely foraging on the same flowers as infected honey bees. I
surveyed honey bee colonies and stingless bee colonies on the campus of the University
of São Paulo in Ribeirão Preto, Brazil and present the first evidence of SBV in colonies
in Brazil; however, I found no evidence of honey bee viruses infecting stingless bees
even in those with colonies near infected honey bees.
iv
Table of Contents
List of Figures .................................................................................................................... vi
List of Tables ................................................................................................................... viii
Acknowledgments............................................................................................................... x
Chapter 1:
1.
Thesis Introduction ....................................................................................... 1
The European Honey Bee, Apis mellifera L. ........................................................... 1
a.
Summary of Honey Bee Biology, Colony Structure, and Lifestyle ....................1
b.
Beekeeping ..........................................................................................................3
2.
Non-Viral Pests and Pathogens of the Honey Bee................................................... 5
3.
Viral Infections of the Honey Bee ........................................................................... 6
a.
Overview of Honey Bee Viruses .........................................................................6
b.
Detection of Honey Bee Viral Infections ............................................................8
Chapter 2:
Exploring the Impact of the Antiviral Drug Ribavirin on RNA Viruses in
Honey Bees ..................................................................................................................... 10
1.
Abstract .................................................................................................................. 10
2.
Introduction ............................................................................................................ 10
3.
Materials and Methods ........................................................................................... 13
4.
Results and discussions .......................................................................................... 18
5.
Acknowledgments.................................................................................................. 49
Chapter 3:
Exploring the Gypsy Moth Caterpillar, Lymantria dispar L., as a System to
Test the Infectivity of a DWV Preparation ....................................................................... 50
1.
Introduction ............................................................................................................ 50
2.
Materials and Methods ........................................................................................... 51
3.
Results and Discussion .......................................................................................... 52
Chapter 4:
First Report of Sacbrood Virus in Honey Bee (Apis mellifera L.) Colonies
in Brazil
..................................................................................................................... 55
v
1.
Abstract .................................................................................................................. 55
2.
Introduction ............................................................................................................ 55
3.
Materials and Methods ........................................................................................... 57
4.
Results and Discussion .......................................................................................... 58
5.
Acknowledgments.................................................................................................. 59
Chapter 5:
Prevalence of Honey Bee Viruses in Native Brazilian Stingless Bees
(Apidae; Meliponini) ........................................................................................................ 60
1.
Introduction ............................................................................................................ 60
2.
Materials and Methods ........................................................................................... 61
3.
Results and Discussion .......................................................................................... 62
Chapter 6:
Conclusion and Summary of results ........................................................... 66
Appendix: Susceptibility of Encapsidated Viruses to RNase Degradation ...................... 68
1.
Materials and Methods ........................................................................................... 68
2.
Results and Discussion .......................................................................................... 69
References ......................................................................................................................... 72
vi
List of Figures
Figure 1: The percent mortality of caged bees fed with 10-ml of sugar water with
different concentrations of cordycepin. Each line represents a different cage with its
corresponding concentration of cordycepin. Percent mortality is the percent of the total
bees that had died at the corresponding time point after treatment began. ........................21
Figure 2: Timeline showing the treatment of ribavirin and other important observations
of the four colonies (T1, T2, C1, and C2) from June 2010 to fall 2011 . Asterisks (*)
indicate the points in time where colonies were sample and tested for viruses. ................25
Figure 3: RT-PCR analysis for actin mRNA on 8 bees per hive collected at four different
times. Treatment colonies were T1 and T2 while control colonies were C1 and C2. ......29
Figure 4: RT-PCR analysis for DWV on 8 bees per hive collected at three different
times. Treatment colonies were T1 and T2 while control colonies were C1 and C2. ......30
Figure 5: RT-PCR analysis for BQCV on 8 bees per hive collected at four different
times. Treatment colonies were T1 and T2 while control colonies were C1 and C2. ......31
Figure 6: RT-PCR analysis for SBV on 8 bees per hive collected at two different times.
Treatment colonies were T1 and T2 while control colonies were C1 and C2. ..................31
Figure 7: RT-PCR analysis for IAPV on 8 bees per hive collected at four different times.
Treatment colonies were T1 and T2 while control colonies were C1 and C2. ..................32
Figure 8: DWV viral titer relative to actin in four colonies at four time points. Treatment
colonies were T1 and T2 while control colonies were C1 and C2. ...................................34
Figure 9: BQCV viral titer relative to actin in four colonies at four time points.
Treatment colonies were T1 and T2 while control colonies were C1 and C2. ..................35
Figure 10: SBV viral titer relative to actin in four colonies at four time points. Treatment
colonies were T1 and T2 while control colonies were C1 and C2. ...................................36
Figure 11: IAPV viral titer relative to actin in four colonies at four time points.
Treatment colonies were T1 and T2 while control colonies were C1 and C2. ..................37
Figure 12: Ion Torrent sequencing coverage across the genome of LSV-2 for RNA
extracted from bees from colony T2 (treated colony) collected June 14, 2010
(pretreatment). ....................................................................................................................42
vii
Figure 13: Ion Torrent sequencing coverage across the genome of DWV for RNA
extracted from bees from colony T2 (treated colony) collected June 14, 2010
(pretreatment). ....................................................................................................................43
Figure 14: Ion Torrent sequencing coverage across the genome of BQCV for RNA
extracted from bees from colony T2 (treated colony) collected June 14, 2010
(pretreatment). ....................................................................................................................44
Figure 15: Ion Torrent sequencing coverage across the genome of BQCV for RNA
extracted from bees from colony T2 (treated colony) collected July, 7 2010. ..................45
Figure 16: The percent of nucleotides differing from the consensus sequence over a 95bp
region of the BQCV genome in the 3’ untranslated region is shown for bees sequenced
from the June 14, 2010 collection of T2 (treated colony). This region from the 8435bp
loci to the 8530bp loci represents the only region with coverage of greater than 1000bp
for the BQCV genome in this sample. ...............................................................................46
Figure 17: DWV RT-PCR assay on RNA from gypsy moths injected with 0.5 µl, 1.0 µl,
or 5.0 µl of either Grace’s media or DWV homogenate in Grace’s media. Gypsy moths
were either frozen immediately (Day 0) or reared for 5 days (Day 5) and then frozen. ....53
Figure 18: DWV positive strand-specific RT-PCR assay on RNA from gypsy moths
injected with 0.5 µl, 1.0 µl, or 5.0 µl of either Grace’s media or DWV homogenate in
Grace’s media. Gypsy moths were either frozen immediately (Day 0) or reared for 5
days (Day 5) and then frozen. ............................................................................................53
Figure 19: DWV negative strand-specific RT-PCR assay on RNA from gypsy moths
injected with 0.5 µl, 1.0 µl, or 5.0 µl of either Grace’s media or DWV homogenate in
Grace’s media. Gypsy moths were either frozen immediately (Day 0) or reared for 5
days (Day 5) and then frozen. ............................................................................................54
Figure 20: Susceptibility of actin mRNA and several honey bee Picornavirles viruses to
ribonuclease A and deoxyribonuclease I digestion. ...........................................................70
viii
List of Tables
Table 1: Summary of the primers used in these experiments for detection of viral
genomes using RT-PCR. ....................................................................................................16
Table 2: Summary of the thermal protocols used to amplify cDNA with viral specific
primers. ..............................................................................................................................16
Table 3: Summary of the primers used for qPCR viral detection. ....................................17
Table 4: Cage bees were fed 10-ml of sugar water with varying concentrations of
cordycepin. The number of dead bees at specific time intervals post-treatment in each
cage is reported here. The remaining number represents the number of live bees left in
each cage 108 hours after the start of treatment. The total number is the number of bees
in each cage at the beginning of the experiment. ...............................................................20
Table 5: Number of dead bees found in ribavirin-treated cages at specific times. Four
cages of approximately 200 bees each were prepared and fed 10-ml of sugar water with
dissolved ribavirin at different concentrations. Live bees were removed at regular
intervals from each cage for testing and the number of dead bees was counted at regular
intervals. .............................................................................................................................22
Table 6: Caged bees were treated with ribavirin in sugar water at varying concentrations
and a sample of five bees from each cage was removed at successive time points. RNA
was extracted from five pooled bees or five individual bees from each cage and
timepoint. RNA was tested for Actin, DWV, and IAPV using a RT-PCR assay. ............23
Table 7: Summary of the feeding and ribavirin treatment schedule over the summer
2010. Four colonies (T1, T2, C1, and C2) were set up in an isolated apiary and given
sugar water with ribavirin (T1 and T2) or plain sugar water (C1 and C2). .......................26
Table 8: Prevalence of RNA viruses in colonies sampled over the course of treatment
with ribavirin. Numbers given indicate how many bees out of eight tested positive for the
virus via virus-specific RT-PCR. Ribavirin-sugar water was given to T1 and T2; C1 and
C2 were only given sugar water.........................................................................................28
Table 9: Summary of PCR products submitted for Ion Torrent sequencing. ...................48
Table 10: Summary of results from RT-PCR analysis for viruses in 10 colonies at the
University of São Paulo in Ribeirão Preto, São Paulo, Brazil. Ten pooled honey bees
ix
from 10 individual colonies were tested for DWV, IAPV, SBV, BQCV, ABPV, CBPV,
and KBV. ...........................................................................................................................62
Table 11: A variety of species of stingless bees from three different meliponiaries were
collected and tested for DWV, IAPV, SBV, and KBV. ....................................................64
Table 12: Concentration of RNA extracted from viral homogenate digested with RNase
A and DNase I for varying amounts of time. RNA was dissolved in a total of 20µl of
RNase free water. ...............................................................................................................70
x
Acknowledgments
I would like to thank Penn State University and the entire Department of Entomology
for the support that they have provided me over the last few years; it has been a pleasure
to work among such great scientists. I would like to especially thank fellow graduate
students Abby Levitt and Rajwinder Singh and laboratory technicians Amanda Fisher and
Monica Kalkstein for invaluable guidance, support, and technical knowledge that they
have provided me.
I would like to the Mr. Jim Bobb for providing the colonies used in the ribavirin field
experiment and Mrs. Harriett Cox for allowing the use of her land for the apiary in Port
Matilda, PA.
The work in Brazil would not have been as successful or enjoyable without the help
and guidance of Dr. David de Jong and numerous other faculty and students at the
genetics department of the University of São Paulo in Ribeirão Preto, Brazil. I would
like to thank them for their hospitality, guidance, and patience while I was there in June
and July 2011.
Funding for this project was provided through the Pennsylvania Pollinator Research
Grant, which was supported by the Pennsylvania Department of Agriculture, the
Pennsylvania Beekeeper’s Association, the Center for Pollinator Research, and the
Montgomery County Beekeepers Association. Funding for travel and living expenses
while I was in Brazil was provided through the Penn State International Program in
Brazil Grant.
I would especially like to thank my advisor Dr. Diana Cox-Foster who assisted in
countless edits of this thesis and has been a tremendous mentor and teacher over the
course of my undergraduate and graduate career at Penn State. Finally, I would like to
thank the additional members of my Master’s committee: Dr. Craig Cameron, Dr. Eddie
Holmes, Dr. Christina Grozinger, and Dr. Ed Rajotte who provided invaluable feedback
and advice over the course of this work.
1
Chapter 1: Thesis Introduction
1. The European Honey Bee, Apis mellifera L.
The western honey bee, Apis mellifera L., is a eusocial insect in the order
Hymenoptera. It is unique as one of the few domesticated insects and is extremely
important to agriculture worldwide. The honey bee produces many commercial products
such as honey, wax, and propolis; however, it is most economically important as a
pollinator and its impact has been estimated to be in excess of $14 billon in the United
States alone (Morse & Calderone, 2000).
In addition to being important to agriculture, the honey bee is one of the most well
studied insects. Much is known about honey bee biology and natural history and its
genome was one of the first insect genomes to be sequenced (The Honeybee Genome
Sequencing Consortium, 2006). This makes the honey bee an ideal model organism for a
variety of studies including those on pathogen ecology and the evolution of social insects.
a. Summary of Honey Bee Biology, Colony Structure, and Lifestyle
A honey bee colony builds its hive in a hollow, enclosed cavity such as a hollow tree
trunk or a man-made hive designed specifically for keeping bees and there are two
primary building materials used: wax and propolis. Wax is secreted from a special gland
on the worker bee and is used to build the comb where honey, pollen, and developing
brood are kept while propolis is a sticky, resinous plant material used as a sealant within
the hive.
One colony can contain up to 60,000 individuals; although, the actual population
varies depending on the time of year and strength of the colony. This superorganism
consists of individuals in one of three different castes: the fertile female gyne (queen), the
sterile female (worker), and the fertile male (drone). These castes can be differentiated
by morphological features and have distinct roles within the colony.
The queen is the only fertile female in the colony. She can be identified by her
elongated, wasp-like abdomen and is about twice the size of the worker bees. A normal
colony has only one and she is responsible for laying all eggs in the colony, up to 2,000
per day. A queen is reared from a normal female egg but is fed royal jelly which contains
2
a specific factor, royalactin, that triggers the development of female larvae into a queen
(Kamakura, 2011). After the virgin queen emerges, she goes on a series of mating flights
to collect semen from drones; a queen can mate with multiple drones with an average of
12 (Tarpy & Nielsen, 2002). Once she returns to the hive and starts laying eggs, she
cannot mate again and must rely on the sperm that she has stored in an organ called the
spermatheca for the duration of her life. With each egg that is laid, she has the option of
fertilizing it so that it will develop into a diploid female or laying an unfertilized egg
which will develop into a haploid male (Mackensen, 1951). The average queen lives
longer than a worker or drone bee, often three to four years, and will be replaced when
the colony recognizes certain signals that she has become unproductive (Butler, 1957).
The worker bees are sterile females and constitute the majority of the total bees in a
colony. Worker bees develop from fertilized, diploid eggs and can live for up to six
weeks in the summer and six months in the winter. Most of the tasks associated with
maintaining a colony are performed by the workers such as: producing wax, caring for
brood, cleaning the hive, defending the hive, storing nectar, feeding and caring for the
queen, and foraging for nectar and pollen. A distinct division of labor among the worker
bees is often observed; however, the role of one individual bee often changes at different
life stages. In general, bees progress over their life cycle from tasks inside the hive such
as rearing brood to tasks outside the hive such as foraging for nectar with older bees
generally performing the latter tasks. This age-based division of labor is, however, highly
variable depending on the needs of the hive and is often difficult to predict (Ribbands,
1952; Rösch, 1925). A worker bee cannot lay fertilized eggs but can lay unfertilized eggs
under certain conditions—typically only in a chronically queenless colony—which
trigger the ovaries of some worker bees to develop allowing them to lay unfertilized eggs
that will ultimately develop into drones.
Drones are haploid males, which develop from unfertilized eggs. They are larger
than worker bees but smaller than queens, and unlike both workers and queens, they lack
a stinger. Their primary purpose is to mate with a queen; although, they do not usually
mate with queens from their own colony. During the early afternoon, drones can be
found in drone congregation areas (Zmarlicki & Morse, 1963) where they are available to
mate with virgin queens. The mating act itself occurs in flight and the drone immediately
3
dies after releasing its semen. The ratio of workers to drones varies throughout the year
as there are more drones in the spring and summer and fewer drones in the winter when it
is less likely that a colony will need to mate a new queen.
All of these individuals function together at the colony level as one superorganism
and rely on a complex array of chemical and visual cues to communicate with each other.
Examples include queen mandibular pheromone (QMP) which is released by the queen
and communicates her presence to the rest of the colony, and the waggle dancing which
allows foragers to communicate the location of a food source to other bees (Frisch, 1967).
Colonies multiply by a process called swarming during which the current queen
leaves the hive with a portion of the workers. The swarm finds a new place to build a
hive and the bees in the original hive rear a new queen. It is unclear exactly what causes
swarms to occur but it is probable that overcrowding and abundance of food are both
important triggers for swarming behavior.
Nectar and pollen are the primary nutrition sources for honey bees. Pollen serves as a
protein source while nectar serves as a carbohydrate source. The primary source of
nutrition of an individual bee differs with the age of the worker and task that the worker
is performing; nurse bees consume more pollen in order to be able to synthesize royal
jelly for the brood (Crailsheim et al., 1992) while foragers consume much more nectar in
order to be able to keep up with the energetic demands of flight. Honey bees also have
several methods of storing food for later use. Bees store nectar in the form of honey by
dehydrating and storing it in a comb with a wax cap. The dehydrated honey is more
energetically dense than nectar and has important antimicrobial properties. Pollen is
stored in the form of bee bread which is a mixture of pollen, honey, and enzymes and is
modified by microorganisms such as yeast (Gilliam, 1979) and lactic acid bacteria
(Vasquez & Olofsson, 2009).
b. Beekeeping
Humans have used honey from honey bees since ancient times. While it is likely that
many hunter-gatherer societies in Europe, Africa, and Asia collected honey from wild
colonies, some of the earliest evidence of the domestication of honey bees comes from
ancient Egyptian tombs and dates as early as the 7th century BC. Initially, honey bees
4
were likely kept in wooden skeps or other hollow containers. In order to extract honey,
the beekeeper removed everything from the hive including the bees, wax, and honey.
The obvious disadvantage of this method was that the colony was killed in this process.
Today, there are several hive designs commonly used which improve upon older designs
but the most common in Europe and the United States is the movable comb hive invented
by Lorenzo Langstroth which he described in his landmark book, The Hive and HoneyBee (1853). His innovative hive design not only allowed the beekeeper to be able to
inspect his colonies but also allowed him to be able to remove honey from the hive
without sacrificing the colony. This transformed beekeeping into a sustainable practice
and made it more feasible to selectively breed honey bees since it was now possible for a
colony to produce honey for the beekeeper and be kept for multiple years.
Because of the numerous benefits of beekeeping, the practice has spread throughout
the world including to the Americas where there were originally no native Apis species
prior to the arrival of Europeans. While the dominant domesticated honey bee worldwide
is the western honey bee, Apis mellifera, the eastern honey bee, Apis cerana F. is
commonly kept in much of Asia. A number of recognized subspecies of both the western
and eastern honey bees are distributed throughout the world, as well, and are often
adapted or have been bred for the specific climate in which they live. An important
example of this is the Africanized honey bee, Apis mellifera scutellata Lepeletier, which
gradually become the dominate species of managed honey bee in South America, Central
America, and parts of North America after it was accidently introduced into southern
Brazil in 1956 (Kerr, 1957). Despite the efforts of beekeepers, the Africanized honeybee
has spread around the tropical Americas, replacing the European subspecies, largely
because of its improved fitness in those climates (Schneider et al., 2004; Spivak, 1992).
Today, honey bees are kept for both honey production and pollination. While honey
is the most visible and direct product of honey bees, the economic value of honey is
dwarfed by honey bee pollination which has been estimated at a value to agriculture in
the United States alone in excess of $14 billion (Morse & Calderone, 2000). Honey bees
are essential pollinators for many crops including almonds, apples, citrus fruit, and alfalfa
and there are many large commercial beekeeping operations that move colonies around
the United States in order to pollinate various crops. One example is the California
5
almond pollination which is the largest managed pollination event in the world and
involves approximately half of all managed honey bee colonies in the United States
(Runckel et al., 2011). Thus, honey bees and the practice of beekeeping are extremely
important to agriculture and their study and preservation is essential not only to the global
economy but also to the global food supply.
2. Non-Viral Pests and Pathogens of the Honey Bee
The health of honey bees is declining around the world and especially in Europe and
North America (Potts et al., 2010; VanEngelsdorp & Meixner, 2010). Although this is
likely caused by numerous factors, pests and pathogens are a definite contributor to this
decline and represent a significant threat to honey bee health. Because of the global
importance of honey bees to agriculture, it is vital to be able to understand and control
these pests.
Honey bee pests and pathogens come from a wide range of taxa and include
vertebrates, arthropods, bacteria, fungi, and viruses. Although the prevalence of specific
pests varies around the world, there are many which are common for honey bees
everywhere. The most problematic vertebrate pests include bears (Ursus species) and
skunks (Mephitis species and Spilogale species) which are capable of completely
destroying a hive to eat the honey and brood inside, and also house mice (Mus musculus
L.) which often seek shelter inside the hive (O’Brien & Marsh, 1990). Important insect
pests include the small hive beetle, Aethina tumida Murray, and the wax moth, Galleria
mellonella L., both of which can destroy the comb structure of a hive. In addition, a
number of insect predators are known to steal honey from honey bee colonies: especially
wasps (Hymenoptera: Vespidae), ants (Hymenoptera: Formicidae), and honey bees from
other colonies. The fugal pathogens Nosema apis and Nosema ceranae can both cause
nonspecific disease symptoms in honey bees and the fungus, Ascosphaera apis, causes
chalkbrood disease which results in dead larvae with a “chalky” appearance. The most
serious bacterial pathogen of honey bees is Paenibacillus larvae which causes American
foulbrood disease.
The most significant parasite of honey bees worldwide and in North America is the
Varroa mite, Varroa destructor (Anderson & Trueman, 2000). This external parasite
6
plagues honey bees by feeding on the hemolymph of prepupae, pupae, and adult bees and
in doing so weakens them and decreases their lifespan. It is also likely that Varroa
infestations can make a colony more susceptible to infection by viral pathogens, both by
weakening the bee’s immune system and by vectoring the virus itself (Shen et al., 2005a,
b; Yang & Cox-Foster, 2005). There are some strains of honey bees which are
particularly resistant to Varroa mite infestation and display “hygienic behavior” where
bees can sense and remove parasitized brood from the colony in order to control an
infestation (Arthi et al., 2000). In addition, it has been proposed that the increased
resistance to Varroa mites shown by Africanized honey bees has contributed to, a least in
part, their success over European honey bees in the tropical Americas (Guzman-Novoa et
al., 1999; Mondragon et al., 2005).
3. Viral Infections of the Honey Bee
a. Overview of Honey Bee Viruses
There are over twenty known viruses that infect honey bees and new viruses are
regularly being discovered (Allen & Ball, 1996; Bromenshenk et al., 2010; Runckel et
al., 2011). Most significant viral pathogens are positive, single-stranded RNA viruses in
the order Picronovirales and family Iflaviridae or Dicistroviridae. Those in the family
Iflaviridae have one open reading frame and include Deformed wing virus (DWV) and
Sacbrood virus (SBV). Those in the family Dicistroviridae have two open reading frames
and include Black queen cell virus (BQCV), Acute bee paralysis virus (ABPV), Kashmir
bee virus (KBV), and Israeli acute paralysis virus (IAPV). An additional RNA virus of
pathological importance is Chronic bee paralysis virus (CBPV) which has a different
genomic structure and remains unclassified (Olivier et al., 2008). Related to CBPV are
another set of RNA viruses called Lake Sinai Virus strains 1 and 2 (LSV-1, LSV-2)
which were recently discovered by deep sequencing (Runckel et al., 2011); although, the
pathologies, if any, associated with these viruses are unknown. Also of note are a family
of DNA viruses invertebrate iridovirus (Iridoviridae), at least one of which is known to
infect honeybees (Bailey et al., 1976).
7
Viruses represent a major threat to honey bee health and cause a variety of
pathologies. The most common honey bee viruses in North America are DWV, SBV,
BQCV, and KBV (Chen & Siede, 2007; Welch et al., 2009). DWV causes bees to
develop with deformed wings unusable for flight. SBV causes Sacbrood disease, a
condition where larvae fail to pupate and die (White, 1913). BQCV is associated with a
blackening of cells containing infected pupae (Bailey & Woods, 1977). KBV does not
have clearly defined symptoms but is associated with honey bee mortality (Chen & Siede,
2007). Recently, viruses have also been implicated as a potential cause of the newly
described honey bee disappearance called colony collapse disorder (CCD) observed in
North America since 2007 (Vanengelsdorp et al., 2009). The newly discovered IAPV
was shown to be strongly correlated with collapsed colonies in a metagenomic study
(Cox-Foster et al., 2007). In addition, an invertebrate iridovirus has been proposed as a
possible cause of CCD (Bromenshenk et al., 2010) although the evidence for this
association remains minimal and disputed (Foster, 2011; Knudsen & Chalkley, 2011;
Tokarz et al., 2011).
A common feature of honey bee RNA viruses and indeed many insect RNA viruses is
that they usually persist as latent, unapparent infections but can replicate rapidly and
cause severe disease under certain conditions such as stress induced by a severe Varroa
mite infection (Martin, 2001). An example of this is DWV which causes honey bees to
develop with ragged wings that are unusable for flight. While this virus is common and
may be detectable year round within a colony, the symptom is most often seen when the
colony is heavily co-infected with Varroa mites during the late summer and fall.
Viruses have been found in bees at all life stages including eggs, larvae, pupae, and
adults and are known to be transmitted in a variety of ways (Chen & Siede, 2007). The
Varroa mite is an important vector and has been shown to be able to transmit DWV,
IAPV, and KBV (Bowen-Walker et al., 1999; Chen et al., 2004; Di Prisco et al., 2011).
In addition to mites, pollen may also be a vector of honey bee viruses (Singh et al., 2010)
providing a way for two insects foraging on the same flower to infect each other. Viruses
can also be transmitted vertically from queens and drones to their offspring (Yue et al.,
2007). Viruses have also been found in pollen, royal jelly, honey (Chen et al., 2006b;
Shen et al., 2005c), and bee feces (Chen et al., 2006a; Hung, 2000) which suggests that
8
they can also be transmitted horizontally between bees in the colony. Inter-taxa
transmission is also possible with some honey bee viruses such as DWV which is also
able to infect and cause disease in bumble bees (Genersch et al., 2006; Singh et al.,
2010).
b. Detection of Honey Bee Viral Infections
While viral infections can sometimes be diagnosed by their symptoms, most
infections remain asymptomatic. This creates a significant challenge for scientists who
study and monitor honey bee viruses. Infections cannot be conclusively diagnosed by the
beekeeper but instead require molecular methods for accurate detection. Broadly, there
are two categories of molecular methods used to detect specific viruses: immunological
methods and genome-based methods. Immunological detection methods use viralspecific antibodies to detect unique epitopes of viral proteins. The most commonly used
immunological assays include: the enzyme-linked immunosorbent assay (ELISA) and
immunodiffusion. Genomic methods, in contrast, derive their specificity by detecting the
unique sequence of the viral genome and include: polymerase chain reaction (PCR),
reverse-transcriptase polymerase chain reaction (RT-PCR), quantitative polymerase chain
reaction (qPCR), microarrays, and viral genomic sequencing.
While immunological methods are still in use today for detection of honey bee
viruses, they have been largely replaced by the use of genomic methods, specifically RTPCR, which is typically faster, less expensive, and more sensitive than immunological
methods (Benjeddou et al., 2001; Chen & Siede, 2007; Genersch, 2005; Grabensteiner et
al., 2001; Shen et al., 2005a). RT-PCR consists of several steps. First, viral RNA must
be extracted from the honey bee. Next, the RNA is converted into a complimentary DNA
(cDNA) library using reverse transcriptase. Finally, the cDNA is amplified by PCR with
viral-specific primers; amplification suggests the presence of the viral genome in the
original RNA sample. Amplification can be detected by electrophoresis of the PCR
product or by monitoring the PCR reaction by real-time PCR (qPCR). Since non-specific
amplification can occur, it is often necessary to sequence the PCR product in order to
confirm that it is virally derived. In these experiments I used RT-PCR and qPCR for
detection of both host-derived and viral-derived RNA.
9
It should also be noted that several controls should be used for the RT-PCR reaction.
A negative control reaction, with no cDNA added, is run to check for contamination in
any of the reagents. A positive control reaction is run by adding a small amount of the
target sequence, in order to verify the viability of the PCR reaction mix and primers. A
PCR reaction with extracted RNA, rather than cDNA, can be run to check for DNA
contamination of the RNA sample. In addition, an endogenous control gene, such as
actin, can be used. Since actin mRNA is transcribed in all honey bee cells, actin can be
amplified in order to verify a successful RNA extraction and cDNA reaction and allows
for the normalization of differences of RNA extractions and cDNA reactions among
different qPCR reactions.
10
Chapter 2: Exploring the Impact of the Antiviral Drug Ribavirin on
RNA Viruses in Honey Bees
1. Abstract
Honey bee viruses represent a significant threat to honey bee health and are
associated with a variety of pathologies such as dead brood, deformed wings, and general
colony mortality. It has also been suggested that Colony Collapse Disorder, the recent
mass-dieoff of honeybees in the North America, is caused by a virus. Despite the threat
posed by honey bee viruses, there are no chemotherapeutic agents currently in use to treat
these viruses and so treatment remains limited to management techniques which are
largely ineffective. Beekeepers are thus often forced to destroy infected colonies in order
to prevent the spread of viral diseases. I explored the broad-spectrum antiviral drug
ribavirin for its potential as an antiviral treatment in honeybees. I present evidence that
ribavirin may be active against Deformed wing virus (DWV), Black queen cell virus
(BQCV), and Sacbrood virus (SBV). I also explored the mutagenic effect of ribavirin on
viral genomes. Although further study is required to determine if ribavirin is safe and can
improve colony survival, this study serves as an important step in the development of an
antiviral treatment for viral infections in honey bees and may also provide insight into the
ecology and epidemiology of these viruses.
2. Introduction
The western honey bee, Apis mellifera L., is an essential pollinator for numerous
crops and has an enormous economic impact (Klein et al., 2007; Morse & Calderone,
2000). Unfortunately, however, honey bee health is declining worldwide (Potts et al.,
2010; VanEngelsdorp & Meixner, 2010). This issue of honey bee decline is complex and
likely has multiple causes however viruses are an important contributing factor because
they are linked to numerous honey bee diseases with a variety of pathologies (Chen &
Siede, 2007). For example, Israeli acute paralysis virus (IAPV) has been associated with
the recent mass die-off of honey bees in North America called colony collapse disorder
(Cox-Foster et al., 2007). Despite the threat posed by viruses, there is little a beekeeper
can do to treat them: treatment is largely limited to management techniques such as the
11
removal of infected brood. Consequently, beekeepers are often forced to destroy
chronically infected colonies to prevent them from infecting others. This practice,
however, limits the productivity and sustainability of beekeeping operations and
increases the overall cost by decreasing yield of honey bee products and pollination
services. Development of an antiviral treatment is therefore necessary and would provide
an essential tool for beekeepers. Some antiviral treatments have been explored such as
Remebee (Beeologics), an RNAi based treatment for IAPV, which is currently
undergoing FDA and EPA trials (Hunter et al., 2010). While RNAi technology shows
promise, it remains to be seen if this can be successfully implemented in the field as a
cost-effective way to control the viruses that plague honey bees. The ideal viral
treatment would be inexpensive, safe, readily available, and active against a wide range
of honey bee viral pathogens. Here I explore the broad-spectrum antiviral drug ribavirin
for the potential to serve as an antiviral treatment in honey bees.
Ribavirin (1-β-D-ribofuranosyl-1,2,4-triazole-3-carboxamide) was chosen for this
study because of its broad-spectrum antiviral properties. It is active against a wide range
of viral taxa including both DNA and RNA viruses (Sidwell et al., 1972). The drug is
used clinically in humans to treat hepatitis C virus (Davis et al., 1998), respiratory
syncytial virus (Wyde, 1998), and lassa fever virus (McCormick et al., 1986).
Promisingly, it is active against foot-and-mouth disease virus (De La Torre et al., 1987),
human rhinovirus, and poliovirus (Anderson et al., 1992), which are all in the same order,
Picornavirales, as many common RNA viruses infecting honey bees. In addition, the
generic status of the drug means that it is relatively inexpensive.
The mechanism of action of ribavirin is best understood for RNA viruses. Ribavirin
is a nucleoside analogue which has considerable structural similarity to purine RNA
nucleotides, a property which allows the drug to interfere with several aspects of RNA
production. The active form of ribavirin is ribavirin triphosphate (RTP) (Streeter et al.,
1973) and the drug is triphosphorylated in the cell by cellular deoxyadenosine kinases
(Streeter et al., 1974). RTP is a powerful competitive inhibitor of the enzyme inosine-5'monophosphate (IMP) dehydrogenase which catalyzes a key step in the production of
guanosine monophosphate (GMP) (Streeter et al., 1973). Consequently, treatment with
ribavirin results in a reduced intercellular concentration of both GMP and its
12
triphosphorylated form, guanosine triphosphate (GTP). A cellular deficiency of GTP can
partially explain the antiviral activity and clinical side effects of ribavirin since GTP is
essential to cellular and viral processes including protein and RNA synthesis. While this
is true, it does not completely explain the antiviral effects of ribavirin (Gilbert & Knight,
1986).
At least four additional mechanisms of action of ribavirin have also been proposed
(Graci & Cameron, 2006) however the most well studied is that of lethal mutagenesis.
Crotty et al. (2000) demonstrated that cellular RTP is incorporated into replicating
poliovirus by the virally encoded RNA-dependent RNA polymerase. The ribavirin base
can pair equally well with either cytidine or uridine. In this way, ribavirin takes the place
of either guanosine or adenosine in the actively replicating viral RNA strand. When the
viral genome with incorporated ribavirin is recopied, either cytosine or uracil forms a
base pair with the ribavirin and is added to the elongating RNA. Thus, C-to-U and G-toA transition mutations are induced by ribavirin (Crotty et al., 2000).
This antiviral strategy of “lethal mutagenesis” relies on the deleterious effects of the
drug-induced mutations (Bull et al., 2007). RNA viruses—which have naturally high
mutation rates—produce fewer viable progeny with the increased mutation rate induced
by the drug. With fewer viable progeny, the infection within a host becomes less severe,
potentially allowing the host to better control or eliminate the infection. If the mutation
rate reaches a certain “extinction threshold” the infection is no longer able to produce
enough viral progeny to sustain itself and the population will go extinct within the host
(Bull et al., 2007).
Lethal mutagenesis is a mechanism particularly relevant to honey bees since the most
important viral pathogens for honey bees are positive, single-stranded RNA viruses in the
order Picornavirales and family Iflaviridae or Dicistroviridae. Those in the family
Iflaviridae have one open reading frame and include Deformed wing virus (DWV) and
Sacbrood virus (SBV). Those in the family Dicistroviridae have two open reading frames
and include Black queen cell virus (BQCV), Acute bee paralysis virus (ABPV), Kashmir
bee virus (KBV), and Israeli acute paralysis virus (IAPV). An additional RNA virus of
pathological importance is Chronic bee paralysis virus (CBPV) which has a different
genomic structure and remains unclassified (Olivier et al., 2008). Related to CBPV are
13
another set of RNA viruses called Lake Sinai Virus strains 1 and 2 (LSV-1, LSV-2)
which were recently discovered by deep sequencing of migratory colonies (Runckel et
al., 2011), although the pathologies associated with these viruses are unknown. The most
common honey bee viruses in North America are DWV, SBV, BQCV, and KBV (Chen
& Siede, 2007; Welch et al., 2009).
In this study, I explore ribavirin for use as a potential antiviral treatment in honey
bees by preforming preliminary tests of the drug on both caged bees and live colonies. I
address several key questions: (i) What is an effective dose of ribavirin? (ii) Is ribavirin
safe for honey bees? (iii) Can ribavirin be used to treat viral infections in honey bees?
(iv) What are the effects of ribavirin on the viral genome? Here, I present preliminary
evidence that ribavirin is active against several important honey bee viruses including
SBV, DWV, and BQCV.
3. Materials and Methods
Collection of Bees for Cage Studies
A sample of worker bees was collected from a queenless colony kept in a greenhouse
which was known to be infected with several viruses. Bees were put into a cold room at
4°C for ten minutes and then separated into mesh cages. Mesh cages with bees were then
put in a dark incubator at 35°C with a pan of water to maintain humidity. Bees were
given an excess of 50% sucrose solution.
Feeding Cordycepin and Ribavirin to Caged Bees
A 10-ml solution of 50% sucrose with varying concentrations of cordycepin, a
nucleoside analogue used as a positive control, or ribavirin was given to each cage with
each cage receiving a different concentration of either cordycepin or ribavirin. Bees were
kept in an incubator with a water pan near the fan to maintain humidity. The incubator
was kept at a constant temperature of 35°C with no interior lighting.
14
Collection of Samples from Caged Bees
Live or dead bees from each cage were collected in a 15 ml sterile screw-top conical
tube at successive time points. Bees were then immediately frozen at -80°C for further
analysis.
Preparation of Colonies
Four honey bee nucleus colonies were prepared in an isolated apiary in Port
Matilda, Centre County, Pennsylvania at N 40° 47.588 W 78° 4.642 during the early
summer 2010. Each of the four colonies was prepared by splitting one parent colony so
that all bees and all four queens were produced from the original colony. Each nucleus
colony was put in two standard five frame deep wooden hive boxes on top of each other.
In this arrangement, the bottom box contained five deep frames while the top box was
empty allowing for the placement of a feeder. The four hives were set next to each other,
about 0.5 meters apart. The hives were each placed on top of individual cinderblocks
with the entrance angled slightly downward and located inside an electric fence in order
to prevent attacks by vertebrate pests such as bears. An index card with a unique design
was placed above the entrance of each colony.
Feeding and Ribavirin Treatment of Colonies
A 50% (v/v) sucrose syrup was prepared by mixing food grade cane sugar (sucrose)
with an equal volume of tap water. The syrup was then poured into a feeder which was
placed inside the hive in the empty box on top of the frames of the box below it. For
colonies which received ribavirin treatment, the appropriate amount of ribavirin was first
dissolved in distilled water and then added to and mixed with the syrup before treating
the hive. Over the course of the summer, two of the colonies were treated with ribavirin
while the remaining control colonies received only sugar water.
Collection of Samples from Colonies
Nurse worker bees were collected by scooping bees from brood comb into sterile 15ml plastic conical screw top tubes. All samples of bees were stored on dry ice until they
were transferred to a -80°C freezer where they were stored for further analysis.
15
RNA Extraction
Bees were sorted on dry ice and homogenized, either individually or in pools, in
500µl of TRIzol reagent (Invitrogen). RNA was extracted in accordance with the
manufacturer’s instructions and suspended in 20µl of RNase-free water (Promega). The
concentration of RNA was measured using a SpectraMax 250 spectrophotometer
(Molecular Devices), a NanoDrop 2000 spectrophotometer (Thermo Scientific), or the
Qubit Fluorometer (Invitrogen) with the RNA protocol.
First-Strand cDNA Library Synthesis
A complimentary DNA library was made from 5µg total RNA using M-MLV reverse
transcriptase (Promega). The cDNA reaction was primed using random primers
(Promega, Cat# C1181).
Viral Specific PCR Reaction
GoTaq green master mix (Promega) was used to amplify cDNA with viral specific
primers. Individual reactions were prepared with 1µl of cDNA and forward and reverse
primers each at a concentration of 0.4µM. Reactions were subjected to one of three
thermal protocols (Table 1) depending on the annealing temperature of the primer.
Primer sequences and corresponding PCR temperature protocol are summarized in
(Table 2).
16
Table 1: Summary of the primers used in these experiments for detection of viral
genomes using RT-PCR.
Virus
IAPV
DWV
DWV
KBV
BQCV
SBV
ABPV
LSV-2
Gene
Product (bp)
Forward (5'-3')
Reverse (5'-3')
Thermal Protocol
Reference
Honeybee Actin
514
ATGAAGATCCTTACAGAAAG
TCTTGTTTAGAGATCCACAT
PCR1
Shen et al. 2005c
Bumble Bee Actin
218
GGAGAAACTTTGTTACGTCGCC
CGCACTTCATGATCGAGTTG
PCR1
Singh et al. 2010
18s Ribosomal RNA
470
GCCAGCGATCCGCCGAAGTT
GCGTGCGGCCCAGAACATCT
PCR3
This Thesis
Singh et al. 2010
Capsid
840
GGTCCAAACCTCGAAATCAA
TTGGTCCGGATGTTAATGGT
PCR2
Singh et al. 2010
Capsid (VP1a)
424
CTCGTCATTTTGTCCCGACT
TGCAAAGATGCTGTCAAACC
PCR2
Capsid (VP1b)
651
GGCGTGGTTCATTAGAATATAGG
AAGCAGATCCCCACCTAAAAA
PCR2
Singh et al. 2011
Capsid
625
TGTTTGTGGCAATCCAGCTA
TACGTCTTCTGCCCATTTCC
PCR3
Singh et al. 2012
Capsid
792
TGGCAACCTAGCCATTTAGC
GGTAGTGGGAGCTGACCAAA
PCR3
Benjeddou et al. 2001
Capsid
210
CACTCAACTTACACAAAAAC
CATTAACTACTCTCACTTTC
PCR1
Shen et al. 2005c
Benjeddou et al. 2001
RNA Polymerase
900
TTATGTGTCCAGAGACTGTATCCA GCTCCTATTGCTCGGTTTTTCGGT
PCR3
Runckel et al. 2011
658
CGGCTGGTCTAGCGTGGCTG
TGGCAAGCTGTGACGAATCCCT
PCR3
Table 2: Summary of the thermal protocols used to amplify cDNA with viral specific
primers.
PCR 1
1 Hold
Temp. (°C)
Time
PCR 2
94
8 min
1 Hold
35 Cycles
1 Hold
94
51.5
72
72
55 sec 55 sec 1:25 sec 10 min
38 Cycles
1 Hold
Temp. (°C)
94
94
55
72
72
Time
8 min
1 min
1 min
1:15 sec 10 min
PCR 3
1 Hold
35 Cycles
1 Hold
Temp. (°C)
94
94
57.5
72
72
Time
8 min
55 sec 55 sec 1:25 sec 10 min
17
Visualization of PCR Product
PCR products were loaded on a 1.5% agarose gel with a 100bp DNA ladder and
separated by electrophoresis.
Viral Specific Quantitative PCR Reaction
TaqMan master mix (Applied Biosystems) was used to amplify cDNA with actin
and viral specific quantitative PCR primers (Table 3). Individual reactions were
prepared with 1µl of cDNA with forward and reverse primers each at a concentration of
0.1µM in a total reaction volume of 20µl. Reactions were run on a 96-well Fast 7500
qPCR machine (Applied Biosystems). The disassociation curves were verified and the Ct
values were calculated using the Fast 7500 Software.
Table 3: Summary of the primers used for qPCR viral detection.
Virus or Gene
Forward (5'-3')
Reverse (5'-3')
Reference
Actin
ATGCCAACACTGTCCTTTCTGG
GACCCACCAATCCATACGGA
Yang et al. 2005
DWV
GACAAAATGACGAGGAGATTGTT CAACTACCTGTAATGTCGTCGTGTT
Yang et al. 2006
IAPV
CGAACTTGGTGACTTGAAGG
GCATCAGTCGTCTTCCAGGT
Cox-Foster et al. 2007
BQCV
GGTGCGGGAGATGATATGGA
GCCGTGTGAGATGCATGAATAC Chantawannakul et al. 2005
Runckel et al. 2011
LSV-2
CGTGCTGAGGCCACGGTTGT
CCGGTGTCGATCTCGCGGAC
Enrichment for RNA Viruses by Filtration and Nuclease Digestion
Six bees were homogenized in a total of 2ml of sterile bee Ringer’s solution
(155mM NaCl, 3mM KCl, 2mM CaCl2). Homogenate was centrifuged and the
supernatant was filtered through a sterile 0.2µm cellulose filter to remove bacteria. Four
hundred microliters of filtrate was digested with 50µl each of RNase A (10mg/ml,
Affymetrix) and DNase I (1mg/ml, Affymetrix) for 45 minutes at 37°C to digest
unencapsidated RNA and genomic DNA. RNA was then extracted from digested filtrate
using 3.5ml of TRIzol reagent (Invitrogen). RNA pellet was resuspended in 19µl of
RNase-free water and 1µl of SUPERase·In RNase inhibitor (Ambion) and stored at 80°C.
Random PCR Amplification of cDNA for Ion Torrent Sequencing
First-strand cDNA synthesis was performed using 2µl of each RNA sample and
Superscript III reverse-transcriptase (Invitrogen) according to the manufacturer’s
18
instructions. The reaction was primed with tagged random septamers at 0.2µM (5’GCCGGAGCTCTGCAGATATCNNNNNNN-3’) and tagged oligo-dT primers at 0.2µM
(5’-GCCGGAGCTCTGCAGATATCNNNNNNN-3’) (Allander et al., 2005; Blomström
et al., 2010). Second-strand cDNA synthesis was performed by incubating the sample at
37°C for 1 hour with 1µl of the Klenow fragment (New England Biolabs).
The cDNA (10µl) was amplified using Platinum pfx DNA polymerase (Invitrogen)
the tag primer (5’-GCCGGAGCTCTGCAGATATC-3’) at a concentration of 0.6µM with
the following thermal protocol: an initial denaturation at 94°C for 8 minutes; 40 cycles of
94°C for 30 seconds, 58°C for 1 minute, 68°C for 2 minutes; a final elongation at 68°C
for 10 minutes.
Ion Torrent Sequencing
Samples were digested with EcoRV (New England Biolabs) and RNase A
(Affymetrix) for 1 hour at 37°C to remove the PCR tag and excess RNA. Digested
samples were purified using the QIAquick PCR purification kit (Qiagen). Samples were
then sent to the Penn State Genomics Core Facility – University Park, PA for library
preparation and Ion Torrent sequencing.
4. Results and discussions
What is an effective dose of ribavirin?
In order to estimate an effective dose of ribavirin, the chain-terminating nucleoside
cordycepin (3’-deoxyadenosine) was given to honey bees as a positive control.
Cordycepin is a toxin originally isolated from Cordyceps, a genus of entomopathogenic
fungi (Cunningham et al., 1950). Cordycepin is triphosphorylated by cellular kinases and
then incorporated into replicating mRNA in place of adenosine by cellular RNA
polymerases. Since cordycepin lacks the 3’-hydroxyl group present on the adenosine
nucleoside, the incorporated cordycepin cannot accept an additional nucleoside and
mRNA synthesis is thus terminated (Müller et al., 1977). While this mechanism of
toxicity in cordycepin is different than the mechanism of action of ribavirin—cordycepin
19
is a chain terminating nucleoside while ribavirin is a mutagen—their mechanisms share a
fundamental similarity. They both work by being improperly incorporated into
replicating RNA. The toxic effect of cordycepin, mortality, is much easier to observe
than the antiviral effects of ribavirin and this feature made cordycepin a good a positive
control to estimate an effective dose of ribavirin. While it is not possible to know exactly
how the dosage and effect ratio compares between ribavirin and cordycepin, this at least
provides a reference point toward estimating an effective dose of ribavirin.
Four cages of approximately 100 bees each were prepared. Each cage was given a
10-ml solution of sugar water with cordycepin at one of four concentrations: 0, 0.4mM,
1.0mM, or 1.3mM. These doses were chosen based on active doses of ribavirin in tissue
culture (Crotty et al., 2001; De La Torre et al., 1987). The mortality in each cage was
measured daily. Comparable mortality was seen in all treatments after a period of 24
hours, suggesting that cordycepin needs at least 24 hours to show an effect. A
considerable difference in mortality between treated and untreated bees was seen after 48
hours with the two highest treatments having mortality rates of 53% and 49% while the
untreated bees had a mortality rate of only 11%. Table 4 summarizes the number of bees
found dead at each time point for each of the four different cordycepin concentrations.
Figure 1 depicts the percent of total bees dead at each time point. By the end of the
experiment, over 90% of bees in the two highest treatments had died while fewer than
50% of the non-treated bees had died. The lowest cordycepin treatment, 0.4mM, also
showed an increase in mortality rate over the non-treatment control, although it was not
as large as the mortality seen in the higher treatments suggesting that toxic levels had not
been reached in this dose. The significant increase in mortality by cordycepin treatment
suggested that ribavirin would also be effective at these dosages and may provide a good
reference concentration for the treatment of honey bees with ribavirin.
20
Table 4: Cage bees were fed 10-ml of sugar water with varying concentrations of
cordycepin. The number of dead bees at specific time intervals post-treatment in each
cage is reported here. The remaining number represents the number of live bees left in
each cage 108 hours after the start of treatment. The total number is the number of bees
in each cage at the beginning of the experiment.
Concentration of Cordycepin (mM)
Time (hours) 0.0
0.4
1.0
1.3
24
4
9
13
5
48
10
11
33
35
72
6
16
25
24
96
7
10
7
11
108
11
9
4
3
Remaining
49
42
5
3
Total
87
97
87
81
21
100%
90%
80%
Percent Mortality
70%
60%
1.3mM
50%
1.0mM
0.4mM
40%
0
30%
20%
10%
0%
20
40
60
80
Time post-treatmtnet (hr)
100
120
Figure 1: The percent mortality of caged bees fed with 10-ml of sugar water with
different concentrations of cordycepin. Each line represents a different cage with its
corresponding concentration of cordycepin. Percent mortality is the percent of the total
bees that had died at the corresponding time point after treatment began.
Is ribavirin safe for honey bees?
The next step of this study was to examine the safety of ribavirin on caged bees. Four
cages of approximately 200 bees each were prepared from bees in a colony known to be
infected with several viruses. After one day in the incubator, each cage was given a 10ml solution of sugar water with dissolved ribavirin at varying concentrations. Three of
the cages were given ribavirin at either 0.4mM, 1.0mM, 1.3mM as experimental groups
and the fourth cage received only sugar water as a negative control. All cages were also
given excess distilled water. After all of the initial sugar water treatment was consumed,
the cages were then given an excess of sugar water without dissolved ribavirin.
22
Five live bees from each cage were collected at successive time points. Each cage
was sampled three times per day for the first two days of the experiment, twice per day
for the next two days of the experiment and then once per day for the next 12 days after
which the experiment was terminated. Dead bees were removed and counted at 54 hours
and 12 days after the beginning of the treatment (Table 5).
Table 5: Number of dead bees found in ribavirin-treated cages at specific times. Four
cages of approximately 200 bees each were prepared and fed 10-ml of sugar water with
dissolved ribavirin at different concentrations. Live bees were removed at regular
intervals from each cage for testing and the number of dead bees was counted at regular
intervals.
Ribavirin (mM)
Time (hours)
0.0
0.4
54 Hrs
6
3
12 Days
54
34
% Mortality 30%
19%
1.0
3
55
29%
1.3
7
67
37%
RNA was extracted from five individual bees from each cage from the zero time point
collection and the 144 hour post-treatment collection. RNA was extracted from five
pooled bees from each cage for each of the 24 hour, 48 hour, 66 hour, and 90 hour posttreatment collections. All RNA extractions were tested by RT-PCR for DWV and IAPV.
No significant difference in health or mortality rates (Table 5) was observed among
the four treatments or controls suggesting that ribavirin is safe at this dosage. All bees or
pools of bees tested positive for DWV while the prevalence of IAPV was minimal and
the prevalence of either virus did not appear to change over the course of the experiment.
(Table 6).
23
Table 6: Caged bees were treated with ribavirin in sugar water at varying concentrations
and a sample of five bees from each cage was removed at successive time points. RNA
was extracted from five pooled bees or five individual bees from each cage and timepoint.
RNA was tested for Actin, DWV, and IAPV using a RT-PCR assay.
[Ribavirin]
(mM)
0.0
0.4
1.0
1.3
∗
Time Post Treatment
(hr.)
Samples
*
% DWVVp1a
DWV-Vp1a
Presence
0
5
100
+
24
1
+
48
1
+
66
1
+
90
1
+
114
5
100
+
0
5
100
+
24
1
+
48
1
+
66
1
+
90
1
+
114
5
100
+
0
5
100
+
24
1
+
48
1
+
66
1
+
90
1
+
114
5
100
+
0
5
100
+
24
1
+
48
1
+
66
1
+
90
1
+
114
5
100
+
Samples were extracted as either 5 individual bees (N=5) or a pool of 5 bees (N=1)
IAPV was also determined for all samples and found to be missing from all samples
except for a single bee at 114 hrs in the 1.3 mm Ribavirin treatment
24
While this result suggested that ribavirin was not effective at clearing a DWV
infection within a honey bee, it does not indicate that the drug is ineffective or useless
against DWV or any other virus. With the additional deleterious mutations induced by
the drug, the virus produces fewer infectious genomes (Bull et al., 2007). Thus, the
question of whether the virus remains infectious in the host becomes more relevant than
whether the virus remains detectable in the host. In honeybees, a treatment which
prevents transmission between individuals in the colony may be sufficient at clearing a
viral infection since horizontal transmission between bees within a colony is an important
viral transmission route and thus probably contributes to sustaining a viral infection
within a colony over successive generations. Even if the ribavirin treatment is capable of
rendering DWV uninfectious without completely clearing the virus from the bees, DWV
may still be detectable with the RT-PCR assay used in this experiment. Thus, the next
step in this study is to treat infected colonies with ribavirin in order to determine whether
the drug has an impact on RNA viruses within a colony.
Can ribavirin be used to treat viral infections in honey bees?
In early June of 2010, four honey bee colonies (T1, T2, C1, C2) were set up in an
isolated apiary in Port Matilda, Centre County, Pennsylvania. Near the apiary was a
small stream and ample floral resources. This apiary was set up away from other honey
bee colonies in order to minimize the potential for ribavirin-contaminated honey to be
robbed by other colonies and also minimize the potential for other colonies to infect the
experimental colonies. To my knowledge, there were no other apiaries within one mile
of this apiary.
Each of the four colonies was prepared by splitting one parent colony so that all bees
and all four queens were progeny of the queen of the parent colony: this was done in
order to minimize genetic diversity among the colonies and to ensure that each colony
had the same initial viruses. An index card with a unique design was placed above the
entrance of each colony so that honey bees would be better able to identify their own
colony in order to minimize drifting.
Two colonies (T1 and T2) were chosen as experimental colonies that were given the
same ribavirin treatment over the summer in the form of ribavirin dissolved in sugar
25
water. The remaining two colonies (C1 and C2) were chosen as control and given only
sugar water without ribavirin. Treatment began on June 30, 2010 and continued until
September 19, 2010 (Figure 2). The dose of ribavirin was chosen based on the amount
of cordycepin that was shown to cause mortality in caged bees. This number was then
extrapolated for the estimated number of bees in each colony. The weekly dose was
increased (Table 7) over the summer as the colonies strengthened. Each of the four
colonies consumed most of the sugar water or sugar water with ribavirin that they were
given and in fact consumed the entire amount of sugar water with ribavirin or plain sugar
water that they were given for every feeding after July 7, 2010 (Table 7). During this
time, the bees were storing honey, so it is possible that the ribavirin-sugar water solution
was stored instead of being immediately consumed.
Figure 2: Timeline showing the treatment of ribavirin and other important observations
of the four colonies (T1, T2, C1, and C2) from June 2010 to fall 2011 . Asterisks (*)
indicate the points in time where colonies were sample and tested for viruses.
26
Table 7: Summary of the feeding and ribavirin treatment schedule over the summer 2010.
Four colonies (T1, T2, C1, and C2) were set up in an isolated apiary and given sugar
water with ribavirin (T1 and T2) or plain sugar water (C1 and C2).
Date
6/10/2010
6/14/2010
6/18/2010
6/30/2010
7/2/2010
7/7/2010
7/9/2010
7/13/2010
7/15/2010
7/20/2010
7/23/2010
7/28/2010
8/2/2010
8/5/2010
8/10/2010
8/15/2010
8/24/2010
8/28/2010
9/5/2010
9/9/2010
9/11/2010
9/19/2010
10/3/2010
Amount of 50% (v/v) Sugar Syrup Given (L) Ribavirin Treatment (mg)
T1
T2
C1
C2
T1
T2
C1
3.8
3.8
3.8
3.8
0
0
0
1.9
1.9
1.9
1.9
0
0
0
1.9
1.9
1.9
1.9
0
0
0
1.0
1.0
1.0
1.0
70
70
0
1.0
1.0
1.0
1.0
70
70
0
1.0
1.0
1.0
1.0
70
70
0
1.0
1.0
1.0
1.0
70
70
0
1.0
1.0
1.0
1.0
70
70
0
3.0
3.0
3.0
3.0
210
210
0
3.0
3.0
3.0
3.0
210
210
0
3.0
3.0
3.0
3.0
210
210
0
3.0
3.0
3.0
3.0
210
210
0
3.0
3.0
3.0
3.0
210
210
0
3.0
3.0
3.0
3.0
840
840
0
3.0
3.0
3.0
3.0
840
840
0
3.0
3.0
3.0
3.0
840
840
0
3.0
3.0
3.0
3.0
840
840
0
3.0
3.0
3.0
3.0
840
840
0
3.0
3.0
3.0
3.0
840
840
0
3.0
3.0
3.0
3.0
840
840
0
3.0
0.0
3.0
0.0
840
0
0
3.0
3.0
3.0
3.0
840
840
0
3.0
3.0
3.0
3.0
0
0
0
C2
0
0
0
0
0
0
0
0
0
0
0
0
0
0
0
0
0
0
0
0
0
0
0
27
Over the course of summer 2010, the four colonies were generally healthy and
flourished, although colonies T1 and C1 were noticeably stronger than T2 and C2. All
colonies were observed to have healthy, laying queens at each collection point throughout
the summer with the exception of C2 which did not have eggs or a queen when it was
checked on September 11, 2010 or September 19, 2010 but was presumably able to
requeen because a queen and eggs were found when it was checked again on November
3, 2010.
The colonies expanded over the course of the summer and when necessary, an
additional 5-frame deep nucleus box was added to the top of the existing one to provide
additional room in which the colony could expand. By the end of the summer, both T1
and C1 had a total of three nucleus boxes with full frames, plus one empty box for the
feeder while T2 and C2 had only two nucleus boxes full with frames plus empty box for
the feeder. At the end of the summer, all four colonies were transferred into ten-frame
deep bottom supers for the winter. Colonies T1 and C1 were transferred into 2 ten-frame
boxes while T2 and C2 were transferred into one.
All four colonies survived the winter and were alive and active in the spring of 2011,
at least until May of that year. The colonies were not monitored closely, fed or given
ribavirin over the summer 2011 and by fall of 2011 there were no bees in T1 while C1,
T2, and C2 were active. The cause of the colony demise in T1 is not known.
A sample of bees was collected weekly over the summer 2010 and several additional
samples were collected in the spring and summer of 2011. Collections from six different
time points were analyzed for viral infections using a reverse-transcriptase PCR (RTPCR) reaction to detect honey bee actin mRNA (Figure 3) and six different honey bee
RNA viruses: DWV(Figure 4), KBV, BQCV(Figure 5), SBV(Figure 6), IAPV(Figure
7), and ABPV. A total of eight bees from each colony and time point were analyzed. A
summary of results is available in Table 8.
28
Table 8: Prevalence of RNA viruses in colonies sampled over the course of treatment
with ribavirin. Numbers given indicate how many bees out of eight tested positive for the
virus via virus-specific RT-PCR. Ribavirin-sugar water was given to T1 and T2; C1 and
C2 were only given sugar water.
VIRUS
DATE
DWV
6/14/10
7/28/10
9/19/10
4/4/11
5/21/11
COLONY
T1
7/8 88%
8/8 100%
8/8 100%
6/8 75%
7/8 88%
BQCV
6/14/10
7/28/10
9/19/10
4/4/11
5/21/11
7/8 88%
8/8 100%
7/8 88%
2/8 25%
5/8 63%
8/8 100%
7/7 100%
7/8 88%
7/8 88%
7/8 88%
8/8 100%
7/8 88%
7/7 100%
2/8 25%
8/8 100%
7/8
6/8
4/8
7/8
7/8
88%
75%
50%
88%
88%
SBV
6/14/10 8/8 100%
4/4/11 1/8 13%
5/21/11 5/8 63%
8/8 100%
7/8 88%
8/8 100%
8/8 100%
2/8 25%
8/8 100%
7/8
0/8
7/8
88%
0%
88%
IAPV
6/14/10
7/28/10
9/19/10
4/4/11
5/21/11
7/8
0/8
0/8
0/8
3/8
88%
0%
0%
0%
38%
8/8 100%
0/7 0%
0/8 0%
0/8 0%
0/8 0%
8/8 100%
0/8 0%
0/7 0%
0/8 0%
2/8 25%
6/8
0/8
0/8
0/8
0/8
75%
0%
0%
0%
0%
KBV
6/14/10 0/8
0%
0/8
0%
1/8
13%
0/8
0%
ABPV
6/14/10 0/8
4/4/11 0/8
5/21/11 0/8
0%
0%
0%
0/8
0/8
0/8
0%
0%
0%
0/8
0/8
0/8
0%
0%
0%
0/8
0/8
0/8
0%
0%
0%
8/8
7/7
8/8
8/8
8/8
C1
100%
100%
100%
100%
100%
8/8
8/8
7/7
8/8
8/8
T2
100%
100%
88%
100%
100%
8/8
8/8
6/8
7/8
6/8
C2
100%
100%
75%
88%
75%
29
Figure 3: RT-PCR analysis for actin mRNA on 8 bees per hive collected at four different
times. Treatment colonies were T1 and T2 while control colonies were C1 and C2.
30
Figure 4: RT-PCR analysis for DWV on 8 bees per hive collected at three different times.
Treatment colonies were T1 and T2 while control colonies were C1 and C2.
31
Figure 5: RT-PCR analysis for BQCV on 8 bees per hive collected at four different times.
Treatment colonies were T1 and T2 while control colonies were C1 and C2.
Figure 6: RT-PCR analysis for SBV on 8 bees per hive collected at two different times.
Treatment colonies were T1 and T2 while control colonies were C1 and C2.
32
Figure 7: RT-PCR analysis for IAPV on 8 bees per hive collected at four different times.
Treatment colonies were T1 and T2 while control colonies were C1 and C2.
33
Next, in order to quantify viral titer, pools of six bees were tested for viral titer
relative to actin using qPCR. Six bees from each colony and each time point were
homogenized together and filtered to remove bacteria. RNA was then extracted from the
viral homogenate and a cDNA library was made. The relative viral titer was then
quantified using primers for Actin, DWV, BQCV, SBV, and IAPV by qPCR (Table 3).
34
10
T1 (Treatment)
T2 (Treatment)
C1 (Control)
C2 (Control)
DWV Viral Titer (2-ΔCt)
1
0.1
0.01
0.001
0.0001
0.00001
0.000001
Collection Date
Figure 8: DWV viral titer relative to actin in four colonies at four time points. Treatment
colonies were T1 and T2 while control colonies were C1 and C2.
35
1
T1 (Treatment)
T2 (Treatment)
C1 (Control)
C2 (Control)
BQCV Viral Titer (2-ΔCt)
0.1
0.01
0.001
0.0001
Collection Date
Figure 9: BQCV viral titer relative to actin in four colonies at four time points.
Treatment colonies were T1 and T2 while control colonies were C1 and C2.
36
1
0.1
0.01
T1 (Treatment)
T2 (Treatment)
C1 (Control)
C2 (Control)
SBV Viral Titer (2-ΔCt)
0.001
0.0001
0.00001
0.000001
0.0000001
1E-08
Collection Date
Figure 10: SBV viral titer relative to actin in four colonies at four time points.
Treatment colonies were T1 and T2 while control colonies were C1 and C2.
37
1
IAPV Viral Titer (2-ΔCt)
0.1
0.01
T1 (Treatment)
T2 (Treatment)
C1 (Control)
C2 (Control)
0.001
0.0001
0.00001
0.000001
0.0000001
Collection Date
Figure 11: IAPV viral titer relative to actin in four colonies at four time points.
Treatment colonies were T1 and T2 while control colonies were C1 and C2.
38
All four colonies were tested for DWV, IAPV, BQCV, SBV, and ABPV from the
pre-treatment collection (June 14, 2011) and had the same prevalence of these viruses at
the beginning of the experiment. They tested positive for DWV, IAPV, BQCV and SBV
(Table 8). Only one colony, T2, had one bee that tested positive for KBV suggesting that
this infection was probably present but only at very low levels. This finding, that all
colonies had the same initial set of viruses at the beginning of the experiment, was
expected since the four colonies were split from one parent colony.
Over the course of the summer, the prevalence of some viral infections did change,
specifically for IAPV, which was detected in all four colonies at the beginning of the
summer but not detected in any of the subsequent collections. While viral prevalence did
change between time points, it remained consistent among the four colonies regardless of
whether they were control colonies or ribavirin treated colonies. Thus, no effect of the
antiviral treatment was measurable using the RT-PCR assay described above for any of
the collections over the summer of 2010.
Interestingly, there was a difference in the prevalence of the virus between the
treatment and control colonies for DWV (Figure 4), BQCV (Figure 5), and SBV
(Figure 6) in bees collected on April 4, 2011. The prevalence and intensity of the viral
bands for these three viruses was greater in the untreated colonies vs. the treated colonies
suggesting that ribavirin had an impact on these viruses. This was also observed in the
RT-qPCR data for SBV (Figure 10) that saw a much greater increase in SBV between
the September 2010 and April 2011 in the untreated colonies compared to the treated
colonies although additional replications will need to be done for this data to be
significant.
It was puzzling, however, that this impact of the drug was not seen until April of
2011, despite the fact that the final treatment with ribavirin was in September of 2010. I
propose several hypotheses for further study to explain this result. First, it may be
possible that while the drug was treating the viruses, treated bees were constantly
becoming re-infected from the environment or other colonies over the course of
treatment. Singh et al. (2010) demonstrated that honey bees can become infected with
RNA viruses simply by foraging on the same flowers as other infected honey bees or
other infected hymenopteran pollinators. Alternately, colonies could have become re-
39
infected through direct contact with infected bees from other colonies such as though
robbing or bees which entered the wrong hive. This is more likely to occur during the
summer than the winter because bees are more active in the summer. Supporting this
hypothesis is the observation that prevalence of these viruses went up in May of 2011
(data not shown). Second, I suggest that while the bees were taking the sugar water with
ribavirin from the feeder, it may not actually have been consumed by the bees but instead
stored for the winter. In this scenario, it was not until the bees began consuming their
winter stores with ribavirin that they were actually receiving the ribavirin treatment. A
deeper understanding of the stability of ribavirin in sugar water will be necessary to
understand the validity of this hypothesis. Additionally, bees in the winter have a longer
lifespan so they would have been exposed to ribavirin for a longer period of time than
summer bees. Finally, I suggest that ribavirin was an effective treatment in these bees
but was unable to completely clear the infections from treated bees but did decrease the
transmissibility of the virus. It was not until new bees were reared in the colony that the
infection was cleared from the colony since the drug treatment prevented the new bees
from becoming infected. An analysis of the sequence variation will provide more insight
into these hypotheses and help to understanding of the impact of ribavirin.
Is ribavirin causing a detectable level of mutations in treated viruses consistent with
the proposed “lethal mutagenesis” mechanism of the drug?
Ribavirin is a mutagen of RNA viruses (Cuevas et al., 2009). To explore this
mechanism of action of ribavirin and determine if ribavirin has an impact in our system, I
examined the intrahost diversity of viral genomes in treated and untreated bees. I
hypothesized that the mutagenic properties of ribavirin will increase the intrahost
diversity of viral sequences within the host. Thus, viral genomes in treated colonies will
show an increase in intrahost diversity of compared to control colonies.
Recent advances in next-generation sequencing (NGS) technology such as 454pyrosequencing, Illumnia, SOLiD, and Ion Torrent provide powerful tools for
determining intrahost diversity. Determining intrahost diversity has previously required
cloning and sequencing many viral genome fragments (Cuevas et al., 2009; Hoelzer et
al., 2010; Murcia et al., 2010); however, it has now been done successfully using NGS
40
(Hoelzer et al., 2010). NGS is faster, more cost-effective per nucleotide sequenced, and
is less labor intensive than traditional cloning and Sanger sequencing. Here I explored
the Ion Torrent PGM platform (Rothberg et al., 2011) of NGS to estimate intrahost
diversity in viral genomes.
I first performed a deep sequencing experiment of the transcriptome. Six bees from
each of four successive collections from T1 (treated colony) were homogenized together
and filtered to remove bacteria. RNA was extracted from each homogenate and a cDNA
library prepared from each RNA sample. Each of the four samples was given a unique
barcode during the library preparation and the samples were run together on the Ion
Torrent PGM sequencer. A total of 4.1 million reads were produced with an average read
length of slightly fewer than 200 base pairs. The four datasets were mapped to the
genomes of DWV, BQCV, IAPV, SBV, LSV-2, CBPV and honeybee ribosomal RNA
using the Bowtie short read aligner (Trapnell et al., 2009) on the Galaxy platform
(Blankenberg et al., 2010; Giardine et al., 2005; Goecks et al., 2010). Only five reads
aligned from one sample with any viral genome, SBV. In fact, the majority of reads were
ribosomal RNA. This suggests that the prevalence of viral RNA relative to ribosomal
RNA was much too low for this deep sequencing method to yield useful results.
Next, I sought to develop a technique to enrich the homogenate for viral RNA so I
explored a method to use RNase A to digest ribosomal RNA. RNase A is an exonuclease
specific to single-stranded RNA. Encapsidated viral RNA should be resistant to RNase A
degradation because the protein capsid can protect the RNA genome from RNase A
degradation. In contrast, free mRNA and ribosomal RNA should be susceptible to
degradation by RNase A. This method has been used successfully for metagenomic
studies using deep sequencing to identify novel viral pathogens (Blomström et al., 2010).
To enrich the preparation for viral RNA, six bees from each of four successive
collections from T2 (treated colony) were homogenized and filtered to remove bacteria.
The filtrate was then digested with RNase A to remove ribosomal RNA and DNase I to
remove DNA. The RNA was extracted from each homogenate and a cDNA library was
prepared from each RNA sample using tagged random and oligo-dT primers for the
cDNA reaction. The cDNA was then amplified using PCR with the tag primer. Each
PCR product was given a unique barcoding sequence during the library preparation and
41
all four samples were run on the Ion Torrent PGM sequencer. The four datasets were
mapped to the genomes of DWV, BQCV, IAPV, SBV, LSV-2, CBPV, and honeybee
ribosomal RNA using the Bowtie short read aligner (Trapnell et al., 2009) on the Galaxy
platform (Blankenberg et al., 2010; Giardine et al., 2005; Goecks et al., 2010). Again,
ribosomal RNA sequences dominated the reads; although, many reads did map to viral
sequences of LSV-2 (Figure 12), DWV (Figure 13) and BQCV (Figure 14 and Figure
15) and the number of reads for each virus reflected the titer of virus present in the bees
at various time points. There was, however, insufficient coverage to use this data to
determine intrahost diversity. For DWV and BQCV, reads tended to be biased toward
the 3’-end of the genome, possibly because of the tagged oligo-dTs used to prime the
cDNA reaction or because of digestion of the 5’-end by the RNase A.
This deep-sequencing experiment did reveal some interesting results. First, it showed
the presence of the recently discovered virus LSV-2 (Runckel et al., 2011) in one of the
samples from T2 collected on June 14, 2010 that I would not have searched for
otherwise. The presence of LSV-2 in this sample was later confirmed by RT-PCR and
qPCR. I then tested all four colonies from collections at four different time points (June
14, 2010; July 7, 2010; September 19, 2010; and April 10, 2011) using RT-PCR and
qPCR; however, LSV-2 was only found in the Jun 14, 2010 collection of T2. Also of
note was a mutation analysis of BQCV. This revealed a region in the 3’-untranslated
region of the viral genome (Leat et al., 2000) from position 8438 to 8444 with a high
incidence of mutations (Figure 16).
42
9000
Sequencing Coverage (Nucleotides)
8000
7000
6000
5000
4000
3000
2000
1000
0
0
1000
2000
3000
4000
Base Position (LSV-2 Partial Genome)
5000
Figure 12: Ion Torrent sequencing coverage across the genome of LSV-2 for RNA
extracted from bees from colony T2 (treated colony) collected June 14, 2010
(pretreatment).
6000
43
140
Sequencing Coverage (Nucleotides)
120
100
80
60
40
20
0
0
2000
4000
6000
8000
Base Position (DWV Genome)
10000
Figure 13: Ion Torrent sequencing coverage across the genome of DWV for RNA
extracted from bees from colony T2 (treated colony) collected June 14, 2010
(pretreatment).
12000
44
5000
Sequencing Coverage (Nucleotides)
4500
4000
3500
3000
2500
2000
1500
1000
500
0
0
1000
2000
3000
4000
5000
6000
Base Position (BQCV Genome)
7000
8000
Figure 14: Ion Torrent sequencing coverage across the genome of BQCV for RNA
extracted from bees from colony T2 (treated colony) collected June 14, 2010
(pretreatment).
9000
45
140
120
Sequencing Coverage (Nucleotides)
100
80
60
40
20
0
0
1000
2000
3000
4000
5000
6000
Base Position (BQCV Genome)
7000
8000
Figure 15: Ion Torrent sequencing coverage across the genome of BQCV for RNA
extracted from bees from colony T2 (treated colony) collected July, 7 2010.
9000
46
4.00%
Total Mutations per Total Coverage
3.50%
3.00%
2.50%
2.00%
1.50%
1.00%
0.50%
0.00%
8420
8440
8460
8480
8500
Base Position (BQCV Genome)
8520
8540
Figure 16: The percent of nucleotides differing from the consensus sequence over a 95bp
region of the BQCV genome in the 3’ untranslated region is shown for bees sequenced
from the June 14, 2010 collection of T2 (treated colony). This region from the 8435bp
loci to the 8530bp loci represents the only region with coverage of greater than 1000bp
for the BQCV genome in this sample.
47
Since neither of the previous techniques provided sufficient data for analysis of
intrahost diversity, I resorted to sequencing specific PCR products amplified from RNA
from homogenized samples. Samples of six bees were homogenized and filtered to
remove bacteria. RNA was extracted and cDNA was prepared using random primers.
The cDNA library was amplified using primers for DWV or BQCV with either GoTaq
green master mix or Platinum Pfx DNA polymerase and purified. Platinum Pfx
polymerase is preferred because it has a smaller error rate compared to GoTaq, however
some samples did not amplify with Platinum Pfx so GoTaq was used instead. PCR
products from both control colonies and both treatment colonies were used so that the
inherent error rate of the sequencing technology could be distinguished from mutations
caused by ribavirin. A total of 32 PCR products were sent for sequencing on two
independent Ion Torrent runs. Table 9 summarizes the samples that were sent for
sequencing. At the time of this writing, sequencing results had been returned by the Penn
State Genomics Core Facility and were being analyzed.
48
Table 9: Summary of PCR products submitted for Ion Torrent sequencing.
Sample Number
Run 1
Run 2
RNA number
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
Colony Collection Date
6/14/2010
346 T1
347 T1
7/7/2010
349 C1
6/14/2010
7/7/2010
350 C1
351 C1
9/19/2010
353 T2
6/14/2010
354 T2
7/7/2010
355 T2
9/11/2010
356 T2
4/10/2011
357 C2
6/14/2010
7/7/2010
358 C2
359 C2
9/11/2010
4/10/2011
352 C1
4/10/2011
360 C2
6/14/2010
357 C2
358 C2
7/7/2010
6/14/2010
345 T1
7/7/2010
346 T1
6/14/2010
353 T2
354 T2
7/7/2010
355 T2
9/11/2010
4/10/2011
356 T2
345 T1
6/14/2010
346 T1
7/7/2010
9/19/2010
347 T1
4/10/2011
348 T1
349 C1
6/14/2010
350 C1
7/7/2010
351 C1
9/19/2010
352 C1
4/10/2011
353 T2
6/14/2010
7/7/2010
354 T2
Primer
DNA Polymerase
DWV-vp1ab
DWV-vp1ab
DWV-vp1ab
DWV-vp1ab
DWV-vp1ab
DWV-vp1ab
DWV-vp1ab
DWV-vp1ab
DWV-vp1ab
DWV-vp1ab
DWV-vp1ab
DWV-vp1ab
DWV-vp1ab
DWV-vp1ab
BQCV
BQCV
BQCV
BQCV
BQCV
BQCV
BQCV
BQCV
BQCV
BQCV
BQCV
BQCV
BQCV
BQCV
BQCV
BQCV
BQCV
BQCV
Platinum pfx
Platinum pfx
Platinum pfx
Platinum pfx
Platinum pfx
Platinum pfx
Platinum pfx
Platinum pfx
Platinum pfx
Platinum pfx
Platinum pfx
Platinum pfx
Platinum pfx
Platinum pfx
GoTaq
GoTaq
Platinum pfx
Platinum pfx
Platinum pfx
Platinum pfx
Platinum pfx
Platinum pfx
GoTaq
GoTaq
GoTaq
GoTaq
GoTaq
GoTaq
GoTaq
GoTaq
GoTaq
GoTaq
Summary of results and ongoing Research
Here, I have explored the impacts of the antiviral drug ribavirin on RNA viruses in
honey bees. I have presented some evidence that ribavirin may be an effective treatment
for honey bee RNA viruses and the results of the most recent sequencing run will help to
elucidate the effect of ribavirin on the genome of replicating RNA viruses. While this
work is an important step in the development of an antiviral drug for honey bees, much
work remains before ribavirin can be used by beekeepers as an antiviral drug. Further
49
research is needed to determine if the drug has an effect on survivorship of the colony,
whether the drug is safe for the environment, and whether there is a risk for the evolution
of ribavirin-resistant viral strains. In addition, the use of ribavirin will probably need to
be strictly limited to colonies which will not produce honey for human consumption since
ribavirin is known to be a teratogen in some animal models (Johnson, 1990) and could
have potentially devastating effects on a developing fetus if it were unintentionally
consumed by a pregnant women.
Before publication of this study, the RNA samples extracted from 8 individual bees
from each of the four colonies at four different time points will be quantified using qPCR.
In addition, the results of pending Ion Torrent sequencing runs will be analyzed.
5. Acknowledgments
I would like to thank Jim Bobb for graciously providing and assisting with the
preparation of colonies used in this experiment and Harriet Cox for the use of her land for
the apiaries in the field studies. I appreciate the help of Deborah Grove and Dan Hannon
at the Penn State Genomics Core Facility - University Park, PA. This work was funded
by the Pennsylvania Pollinator Research Grant, which was supported by the Pennsylvania
Department of Agriculture, the Pennsylvania State Beekeepers Association, the
Montgomery County Beekeepers’ Association, and the Center for Pollinator Research.
50
Chapter 3: Exploring the Gypsy Moth Caterpillar, Lymantria dispar L.,
as a System to Test the Infectivity of a DWV Preparation
1. Introduction
Results of the cage study in the previous chapter suggested that ribavirin was
ineffective at completely clearing an infection of DWV since it was still detectable in
treated bees. Under the lethal mutagenesis model for ribavirin, the drug causes mutations
in the replicating virus. With the additional deleterious mutations induced by the drug,
the virus is unable to produce as many infectious genomes (Bull et al., 2007). Thus, the
question of whether the virus remains infectious in the host becomes potentially more
relevant than whether the virus remains detectable in the host. In honeybees a treatment
which makes the virus less virulent and pathogenic or less transmissible may potentially
be sufficient since bee to bee transmission within the colony may be an important viral
transmission mechanism and thus potentially contributes to sustaining a viral infection
within a colony.
The goal of this experiment was to design and investigate a system to test a DWV
preparation for its infectivity. It was necessary to select a non-honey bee system since
neither DWV-free bees nor honey bee tissue culture cell lines are available and in this
study, I explore the gypsy moth caterpillar, Lymantria dispar L. (Lepidoptera;
Lymantriidae), as a system to test the infectivity of a DWV preparation. The gypsy moth
caterpillar was a convenient system to try because it was readily available in our
department. Although DWV is a honey bee virus, it has been detected in other insects
(Genersch et al., 2006; Singh et al., 2010). In addition, previous studies from our lab
provided some evidence that DWV is capable of infecting gypsy moth cells in tissue
culture; however, from this study I conclude that DWV does not replicate in intact
caterpillars.
51
2. Materials and Methods
Preparation of DWV homogenate
Four bees were cut in half on dry ice; one half was used to confirm the presence of
DWV using RT-PCR while the other half was stored at -80°C for use in preparation of
the homogenate. The bee halves were homogenized in 2 ml of Grace’s insect media
(Sigma), passed through a sterile, 0.2 µm filter, and frozen at -80°C.
Injection of gypsy moth caterpillars
Newly molted fourth-instar Lymantria dispar L. larva were injected with either 0.5
µl, 1.0 µl or 5.0 µl of either Grace’s insect media or the DWV homogenate. Injections
were performed under a stereo dissecting microscope with a sterile 32-guage sharp needle
which was inserted at the base of the second proleg. Four caterpillars were injected with
each combination of volume and either grace’s media or DWV homogenate; two
caterpillars from each combination were immediately frozen at -80°C while the
remaining two were reared for five days.
Rearing of injected caterpillars
The remaining caterpillars, two from each treatment combination, were each put in an
individual cup with artificial gypsy moth diet and incubated at 37°C for five days. After
five days, caterpillars were frozen at -80°C for further analysis.
RNA extraction
RNA was extracted from each individual caterpillar by grinding the sample in 500µl
of TRIzol reagent (Invitrogen) and extracting the RNA in accordance with the
manufacturer’s instructions.
RNA library and Strand-specific cDNA synthesis
Three different sets of complimentary DNA from each extraction were each made
from 5µg total RNA using M-MLV reverse transcriptase (Promega). One set of cDNA
reactions was primed using random primers (Promega). The two remaining sets of
52
cDNA reactions were primed using either forward or reverse DWV primers (Table 1) at
a concentration of 0.75µM each.
DWV specific PCR reaction
GoTaq green master mix (Promega) was used to amplify each of the three cDNA with
primers for DWV. Individual reactions were prepared with 1µl of cDNA and forward
and reverse primers each at a concentration of 0.4µM. Thermal protocols are
summarized in (Table 2). Primer sequences and corresponding thermal protocol are
summarized in (Table 2).
Visualization of PCR product
PCR products were loaded on a 1.5% agarose gel with a 100bp DNA ladder and
separated by electrophoresis.
3. Results and Discussion
The concept behind this technique is based on the ability to detect the anti-sense
replicative intermediate of DWV. This is an indicator of viral replication since an
encapsidated virion carries the positive-sense genome; however, only an actively
replicating infection has the negative-strand replicative intermediate. If just DWV
virions are present, then only the positive strand will be detectable in a strand-specific
RT-PCR assay. If, however, the virus is actively replicating then both the positive and
negative-sense strands will be detectable (Boncristiani et al., 2009).
Gypsy moth caterpillars were injected with varying amounts of either Grace’s insect
medium (control) or a DWV homogenate (experimental). Half of the caterpillars were
frozen immediately while the other half were reared for five days and then frozen. All
gypsy moth caterpillars that were not frozen immediately after injection survived for the
entire five days before they were frozen. A strand-specific RT-PCR assay was used to
detect the positive and negative strand of DWV in each caterpillar. Complimentary DNA
primed with random primers was used to detect the presence of DWV in the gypsy moths
(Figure 17). Complimentary DNA primed with the DWV reverse primer was used to
53
detect the positive strand of the virus (Figure 18). Complimentary DNA primed with the
DWV forward primer was used to detect the negative strand of the virus (Figure 19).
Figure 17: DWV RT-PCR assay on RNA from gypsy moths injected with 0.5 µl, 1.0 µl, or
5.0 µl of either Grace’s media or DWV homogenate in Grace’s media. Gypsy moths
were either frozen immediately (Day 0) or reared for 5 days (Day 5) and then frozen.
Figure 18: DWV positive strand-specific RT-PCR assay on RNA from gypsy moths
injected with 0.5 µl, 1.0 µl, or 5.0 µl of either Grace’s media or DWV homogenate in
Grace’s media. Gypsy moths were either frozen immediately (Day 0) or reared for 5
days (Day 5) and then frozen.
54
Figure 19: DWV negative strand-specific RT-PCR assay on RNA from gypsy moths
injected with 0.5 µl, 1.0 µl, or 5.0 µl of either Grace’s media or DWV homogenate in
Grace’s media. Gypsy moths were either frozen immediately (Day 0) or reared for 5
days (Day 5) and then frozen.
The positive strand and negative strand of DWV were both present in caterpillars
injected with the viral preparation and immediately frozen. This indicates that the virus
was actively replicating in the honey bees used to make the viral preparation. After the
five day incubation period, however, only the positive strand was detectable suggesting
that DWV was probably not actively replicating in this system. Since the encapsidated
positive-sense genome is much more resistant to degradation by host RNases than the
free negative-strand replicative intermediate, it is much more likely that the positive
strand would survive the five day incubation period within the host than the negative
strand. Since only the positive strand was detectable after the five day incubation period,
it is likely that DWV was not actively replicating in this system. This suggests that DWV
was not capable of replicating to high levels over extended time in intact gypsy moths,
although it cannot be ruled out that the homogenate itself was not infectious, perhaps due
to changes in the virus induced by the freeze-thaw cycle. In addition, the initial period
following infection needs to be evaluated to determine if the virus is replicating in the
first few hours. Further experiments could include using a fresh homogenate which was
not frozen and in which only encapsidated virus is injected. In addition, this experiment
could be repeated on gypsy moth cell culture lines which may lack the host defenses of
the caterpillar.
55
Chapter 4: First Report of Sacbrood Virus in Honey Bee (Apis mellifera
L.) Colonies in Brazil
1. Abstract
Sacbrood disease is an affliction of honey bees (Apis mellifera L.) characterized by
diseased brood which fails to pupate and dies and this disease represents a significant
health threat to honey bee colonies. The disease is caused by Sacbrood virus (SBV), a
positive, single-stranded RNA virus in the order Picornavirales which has been found
across the world and recently in some countries in South America. I report the first
evidence of SBV in honey bee colonies in Brazil. SBV was detected using ReverseTranscriptase Polymerase Chain Reaction (RT-PCR) and confirmed with Sanger
sequencing to verify the identity of the PCR product.
2. Introduction
The honey bee, Apis mellifera L., is extremely important to agriculture worldwide,
not only for its honey production but also for its vital role as a pollinator. In Brazil,
honey bee health is declining (Teixeira et al., 2008) and although this is mild compared
to the recent die-off’s reported in Europe and North America (Vanengelsdorp et al.,
2009) it is still of great concern considering the economic importance of honey bees to
Brazilian agriculture. This difference may be related to the fact that the African
subspecies, Apis mellifera scutellata, which was introduced to Brazil in 1956 (Kerr,
1957) is now the dominant type of honey bee found in South America. The Africanized
subspecies seems to show an increased resistance to varroa mites (Guzman-Novoa et al.,
1999; Mondragon et al., 2005).
Honey bee health is influenced by numerous factors including parasites, pathogens,
pesticides, and nutrition. Viral pathogens are often associated with disease and have even
been associated with the recent honey bee die-offs observed in the United States (CoxFoster et al., 2007).
At least 22 different viruses are known to infect the honey bee with the best studied
being positive sense single-stranded RNA ((+)ssRNA) viruses in the order Picornavirales
(Chen & Siede, 2007; Runckel et al., 2011). While many of these viruses can cause
56
disease, they frequently exist in the colony as latent infections that can multiply rapidly
under certain conditions such as when the colony is compromised by some other stress
such as an infestation with the Varroa mite, Varroa destructor Anderson & Trueman,
which can spread the virus (Genersch & Aubert, 2010) and impair the host immune
function (Yang & Cox-Foster, 2005). This presents an additional challenge to the
diagnosis and monitoring of RNA viral infections in honey bees because molecular
techniques are thus required to confirm the presence of the virus rather than visible
symptoms observable by the beekeeper. In the past, RNA viruses were detected by
immunological methods; however, Reverse-Transcriptase Polymerase Chain Reaction
has largely replaced immunological methods as the primary way to detect RNA viruses in
honey bees because of improved accuracy and sensitivity as well as economic benefits
(Benjeddou et al., 2001; Chen & Siede, 2007; Genersch, 2005; Grabensteiner et al.,
2001; Shen et al., 2005a).
Many Picornavirales viruses are known to cause disease in honey bees including the
Iflavirus Sacbrood Virus (SBV), which is the etiological agent of Sacbrood disease in A.
mellifera (Bailey et al., 1964). Sacbrood disease is an important malady of honey bees
and is characterized by larvae which fail to pupate and turn pale yellow and eventually
dark brown before they die. The virus was first isolated and characterized by Bailey et al.
(1964) although the disease itself was first described in 1913 (White) and attributed to a
viral infection in 1917 (White). Since the disease was first characterized, it has been
found throughout Europe, Africa, Asia, North America, and recently South America
where it has been found in both Uruguay(Antúnez et al., 2006) and Argentina (Reynaldi
et al., 2010).
I report the detection, using RT-PCR, of SBV in managed A. mellifera colonies in the
state of São Paulo, Brazil. While a number of other Picornavirales have been found in
colonies in Brazil including Deformed wing virus (DWV), Acute bee paralysis virus
(ABPV), and Black queen cell virus (BQCV) (Teixeira et al., 2008), this represents the
first detection of SBV in Brazilian A. mellifera colonies.
57
3. Materials and Methods
Collection of Bees
All bees were collected from apiaries on the University of São Paulo campus in
Ribeirão Preto, Brazil. Foraging workers were sampled by collection of bees entering the
hive while nurse bees were sampled by collection of bees directly on the brood. Brood
was sampled by careful removal with tweezers. Samples were stored at -80°C until being
processed.
RNA Extraction
RNA was then extracted using the TRIzol reagent (Invitrogen) according to the
manufacturer’s directions. Honey bee RNA was extracted from a pool of 10 bees
homogenized together, from individual bees, or from individual brood. The RNA pellet
was precipitated with isopropyl alcohol overnight at -20°C. The pellet was then washed
with 75% ethanol, resuspended in 20µl of nuclease-free water and stored at -80°C for
further analysis. The concentration of RNA in each sample was measured using a
spectrophotometer (NanoDrop ND-1000).
Reverse Transcriptase PCR
First-strand reverse-transcriptase cDNA synthesis was performed on extracted RNA
samples using 5µg of total RNA and superscript II (Invitrogen) according to the
manufacturer’s instructions. The RNA template was primed using an oligonucliotidedeoxythymidine (oligo-dT) primer.
Viral Specific PCR
Honey bee cDNA was amplified with SBV primers (SBV-F 5’CACTCAACTTACACAAAAAC-3’; SBV-R 5’-CATTAACTACTCTCACTTTC-3’)
specific to a 210bp section in the capsid region of SBV as described by Shen et al. (Shen
et al., 2005c). Samples were amplified with Taq DNA polymerase (Promega) with each
primer at a concentration of 0.4µM. Reactions were carried out on a thermal protocol
consisting of an initial incubation at 94°C for 8 minutes followed by 35 cycles of 94°C
58
for 55 seconds, 51.5°C for 55 seconds, 72°C for 85 seconds with a final incubation at
72°C for 10 minutes.
Gel Electrophoresis
DNA samples were visualized on a 1.5% agarose gel and compared to a standard
100bp molecular weight DNA ladder.
Sequencing
PCR products were precipitated with isopropyl alcohol and resuspended in 10µl of
nuclease-free water. Samples were prepared for sequencing using the BigDye
Terminator v3.1 Cycle Sequencing Kit (Applied Biosystems) and a GeneAmp PCR
System 9600 (Applied Biosystems) in accordance with the manufacturer’s instructions.
Samples were then sequenced using an ABI Prism 310 Genetic Analyzer (Applied
Biosystems).
4. Results and Discussion
An initial survey of bees from 10 colonies in an apiary on the University of São Paulo
campus in Ribeirão Preto was done by pooling 10 forager bees from each colony and
testing them for SBV using the RT-PCR assay described above. Of the 10 colonies
surveyed, one had a positive band of approximately 210bp, suggesting the presence of
SBV in this colony. This colony was retested for SBV using the RT-PCR assay on a total
of 8 individual forager bees; 3 of the 8 bees tested using the RT-PCR assay were positive
for SBV. While the initial colony with SBV was observed to be healthy, an additional
colony was tested in a nearby apiary on the campus that was exhibiting symptoms
suggestive of SBV. From this colony, eight larvae and eight nurse bees were tested for
SBV using the RT-PCR assay described above. Of the larvae tested, 1 of 8 tested
positive for SBV. Of the brood nurse bees tested, 4 of 8 tested positive for SBV. The
PCR product of each reaction was sequenced and aligned to the SBV complete genome
(GenBank accession number AF092924.1) (Ghosh et al., 1999).
This represents the first detection of SBV in Brazil, although it is not surprising that
SBV has been found there considering its worldwide incidence and the recent findings in
59
neighboring countries (Antunez et al., 2006). With global trade and travel occurring at
unprecedented rates, the global spread of viruses and other pathogens can occur faster
than ever before. It is thus extremely important to monitor the global spread of pathogens
including those that infect honey bees. For SBV in particular, further and more extensive
studies will be vital to understanding how the virus spreads as well as the prevalence and
impact of the virus around Southern Brazil and throughout the entire country.
5. Acknowledgments
I would like to thank the Penn State International Graduate Program in Brazil for
providing travel expenses and Dra. Zilá Simões at the University of São Paulo, Ribeirão
Preto campus for providing supplies to do the molecular analysis.
60
Chapter 5: Prevalence of Honey Bee Viruses in Native Brazilian
Stingless Bees (Apidae; Meliponini)
1. Introduction
Stingless bees (Apidae; Meliponini) are a tribe of eusocial bees found throughout the
tropics including South and Central America, Australia, Africa, and Asia. There are over
500 species and many are important natural pollinators for both wild fauna and
agricultural crops (Slaa et al., 2006). In Brazil and other countries, many techniques of
meliponiculture, the management of native bees, have been developed in order to use
native bees as managed pollinators (Roubic et al., 1987). One advantage of keeping
stingless bees is that they lack stingers, making them easier to manage, although many
species display other defense mechanisms such as flying toward and intruder or biting it
and injecting formic acid (Roubic et al., 1987). In addition, several species of stingless
bees can produce a small amount of honey which can be sold for a premium in Brazil and
around the world because of its unique taste and suggested medicinal properties.
There have been a few diseases reported in stingless bees (Nogueira-Neto, 1997);
however, they are poorly studied and almost nothing is known about their causes (NunesSilva et al., 2009). One potential source of diseases in stingless bees is from honey bees.
In North America, some viruses traditionally characterized as honey bee viruses are
known to infect species of native bumble bees and can even cause pathology similar to
that caused by these viruses in honey bees (Genersch et al., 2006). Viruses can also be
transferred between bee species through pollen (Singh et al., 2010) putting native bees at
risk for honey bee viruses when they forage on the same flowers as honey bees. This is
an additional unintended impact that this non-native species may have on native bees.
It is unknown whether honey bee viruses can also infect stingless bees, but it may be
possible considering the inter-taxa range already documented in honey bee viruses.
Given that stingless bees are important insects for both natural and agricultural
ecosystems, it is necessary to explore the potential impact that honeybee viruses may
have on stingless bees. In this experiment, I search for viruses in managed stingless bee
colonies at three sites—which varied in their proximity to managed honey bee colonies—
in the State of São Paulo, Brazil.
61
2. Materials and Methods
Collection of Bees
Foraging worker honey bees and foraging native bees were sampled by collecting
bees entering the hive. Samples were stored at -80°C until being processed.
RNA Extraction
RNA was extracted using the TRIzol reagent (Invitrogen) according to the
manufacturer’s instructions. The RNA pellet was precipitated with isopropyl alcohol
overnight at -20°C. The pellet was then washed with 75% ethanol, resuspended in 20µl
of nuclease-free water and stored at -80°C for further analysis. The concentration of
RNA in each sample was measured using a NanoDrop 1000 spectrophotometer (Thermo
Scientific).
Reverse Transcriptase PCR
First-strand reverse-transcriptase cDNA synthesis was performed on extracted RNA
samples using 5µg of total RNA and superscript II (Invitrogen) according to the
manufacturer’s instructions. The RNA template was primed using an oligonucliotidedeoxythymidine (oligo-dT) primer.
Viral Specific PCR
One microliter of cDNA used for each PCR reaction. Samples were amplified with
Taq DNA polymerase (Promega) with each primer at a concentration of 0.4µM. Viral
specific primers are summarized in Table 1 and corresponding thermal protocols are
summarized in Table 2.
Gel Electrophoresis
DNA samples were visualized on a 1.5% agarose gel and compared to a standard 100bp
molecular weight DNA ladder.
62
3. Results and Discussion
Ten honeybee colonies at the University of São Paulo in Ribeirão Preto, São Paulo,
Brazil were tested for a variety of honey bee viruses in order to first determine what
viruses might be present in honey bee colonies on the campus. A sample of ten forager
bees was collected from ten different colonies in the apiary on campus. RNA was
extracted for the ten bees together and tested using the RT-PCR assay for DWV, IAPV,
SBV, BQCV ABPV, CBPV, and KBV. Of the ten colonies, all tested positive for DWV,
one tested positive for IAPV, one tested positive for SBV, and three tested positive for
KBV. Results are summarized in Table 10.
Table 10: Summary of results from RT-PCR analysis for viruses in 10 colonies at the
University of São Paulo in Ribeirão Preto, São Paulo, Brazil. Ten pooled honey bees
from 10 individual colonies were tested for DWV, IAPV, SBV, BQCV, ABPV, CBPV, and
KBV.
Colony
1
2
3
4
5
6
7
8
9
10
Genus
Apis
Apis
Apis
Apis
Apis
Apis
Apis
Apis
Apis
Apis
Species Actin
mellifera
+
mellifera
+
mellifera
+
mellifera
+
mellifera
+
mellifera
+
mellifera
+
mellifera
+
mellifera
+
mellifera
+
DWV-vp1a IAPV SBV BQCV ABPV CBPV KBV
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
Since honey bees may be a source of viruses in native bees, three meliponiaries
(places with managed stingless bee colonies) were chosen based on their proximity to
honey bee colonies. Meliponiary 1 was located on the campus directly adjacent to the
apiary where the original tested honeybees were kept. Meliponiary 2 was a located
approximately 500m from the original apiary on campus and was separated by a forests
and a lake 100m wide. Meliponiary 3 was a on a farm in São Simão, São Paulo, Brazil
with no managed honeybee colonies within foraging distance. A sample of ten bees was
collected from ten to twelve different colonies in each meliponiary; genus and species of
63
bees are summarized in Table 11. RNA was extracted for the ten bees together and
tested for DWV, IAPV, SBV, and KBV using the RT-PCR assay; these viruses were
chosen because they were first found in the tested honey bee colonies.
64
Table 11: A variety of species of stingless bees from three different meliponiaries were
collected and tested for DWV, IAPV, SBV, and KBV.
Colony Number
Meliponiary 1
1
2
3
4
5
6
7
8
9
10
Meliponiary 2
11
12
13
14
15
16
17
18
19
20
Meliponiary 3
21
22
23
24
25
26
27
28
29
30
31
32
Genus
Tetragona
Tetragona
Frieseomelitta
Scaptotrigona
Partamona
Melipona
Melipona
Nannotrigona
Leurotrigona
Plebeia
Melipona
Melipona
Scaptotrigona
Scaptotrigona
Nannotrigona
Plebeia
Tetragonisca
Tetragona
Partamona
Frieseomelitta
Frieseomelitta
Plebeia
Marmelita
Scaptotrigona
Scaptotrigona
Tetragonisca
Frieseomelitta
Melipona
Melipona
Melipona
Nannotrigona
Scaptotrigona
Species
clavipes
angustula
varia
bipunctata
helleri
quadrifasciata
marginata
testaceicornis
muelleri
remota
scutellaris
quadrifasciata
bipunctata
depilis
testaceicornis
droryana
angustula
clavipes
helleri
varia
polistica
polysticta
angustula
preta
scutellaris
quadrifasciata
testaceicornis
-
65
These native bees were tested for honey bee viruses and no evidence of these viruses
was found in any colonies. While this preliminary result is hopeful, more work needs to
be done to understand whether honey bee viruses or other pathogens are infecting native
bees. This study has several limitations that should be addressed in future studies. First,
the sampling was severely limited in scope. Future studies should involve sampling from
more meliponiaries and wild stingless bee colonies from across Brazil and in other
countries were stingless bees are found. Second, the RT-PCR assay used to detect
viruses is limiting. It is unknown what the minimum infection threshold is for detecting
viruses is, but it is possible that a non-severe infection could potentially go undetected
with the RT-PCR assay. Also, since the RT-PCR assay is sequence-specific, mutations in
the viral genome—which may have been necessary for the cross-species transmission of
viruses—could make the genome undetectable using the same RT-PCR primers if the
mutations occurred at sites where the PCR primers normally bind.
66
Chapter 6: Conclusion and Summary of results
Unfortunately, honey bee health is threatened. Both managed and wild colonies are
dying in many places around the word, especially in North America and Europe (Potts et
al., 2010). Although it is not clear what is causing this decline, it is likely due to a
variety of factors which may include: pesticides, pathogens, human activities, and
management techniques. More research will be necessary to understand this problem and
to work toward a solution to help save honey bees.
One cause of honey bee disease is from a variety of single-stranded RNA viruses in
the order Picornavirales. These can cause a variety of diseases ranging from specific
symptoms such as deformed wings to nonspecific honey bee mortality. Because of the
threat of viruses, the development of a treatment for honey bee viruses has the potential
to help beekeepers around the world. In addition, the lack of a virus-free bee makes the
study of viral transmission in honey bees difficult; a drug capable of clearing viral
infections in honey bees could help researchers who study these viruses.
In this thesis, I explored the drug ribavirin as a potential antiviral agent in honeybees.
I first tested the drug by feeding it to caged honey bees and find evidence to suggest that
ribavirin does not induce mortality in honey bees at potentially therapeutic doses. I next
tested the drug on actual colonies by feeding the drug to bees in a sugar water solution. I
found that ribavirin does seem to have an impact on viral prevalence; although, this
impact was not detectible until the spring after bees were treated with ribavirin. Finally, I
explored several techniques to determine the impact of ribavirin on the intrahost diversity
of viral genomes within honey bees.
While these results are promising, it will be a long time before ribavirin can be used
as a chemotherapeutic agent by beekeepers. A myriad of questions remain and further
studies should focus on whether drug treatment has an effect on survivorship of the
colony, whether the drug treatment poses a risk of contaminating honey or other bee
products that may be consumed by humans, whether the drug is safe for the environment,
and whether the evolution of ribavirin-resistant viral strains is possible. Given that
ribavirin is a teratogen in some animal models and probably humans as well, it is unlikely
that it would be approved for use by beekeepers; other antivirals may offer greater safety.
67
For researchers who can maintain isolated colonies or cages of bees, this treatment does
offer the promise of being able to eliminate virus from those bees to allow for virus-free
bees.
In the next part of this thesis, I searched for honey bee viruses in managed stingless
bee colonies in Brazil. Transmission of honey bee viruses to other hymenopteran
pollinators has been documented and represents an unintended negative consequence to
native pollinators from honey bees which are not native to the Americas. While I found
no evidence of honey bee viruses in the native bees that I tested, a more in-depth study is
necessary to fully understand what risk honey bee viruses have on native stingless bees in
Brazil and throughout the tropics.
Honey bees are unique because they are one of very few insects that have been
successfully domesticated. Humans have used honey from honey bees for thousands of
years because it has traditionally been one of the only available sweeteners. In recent
times, the role of honey bees as managed pollinators has become vital to agriculture and
the global food supply because they are often only insect capable of effectively
pollinating large-scale monocultures common to modern agriculture in developed
countries. The impact of honey bees is evident in many aspects of culture around the
world from simple works of art to multi-million dollar movies (Hickner & Smith, 2007).
In short, honey bees are extremely important to humans. Thus, research that helps to
understand and help honey bee health is extremely important and worthwhile. It is my
sincere hope that this work has contributed to our understanding of viruses in honey bees
and can be developed further to provide additional tools to help save this invaluable
insect.
68
Appendix: Susceptibility of Encapsidated Viruses to RNase Degradation
1. Materials and Methods
Preparation of Viral Homogenate
Eighteen bees were homogenized in a total of 6mL of sterile Bee Ringer’s Solution
(155mM NaCl, 3mM KCl, 2mM CaCl2). Bees used were taken from a colony known to
be infected with several viruses and were stored at -80°C prior to use in this experiment.
Homogenate was centrifuged and the supernatant was filtered through a sterile 0.2µm
cellulose filter.
Nuclease Digestion
Five aliquots of 400µl filtered homogenate were prepared. TRIzol reagent
(Invitrogen) (2.5ml) was immediately added to the zero time point. Fifty microliters each
of RNase A (10mg/ml, Affymetrix) and DNase I (1mg/ml, Affymetrix) were added to
each aliquot. Each aliquot was incubated at 37°C for a varying amount of time: 15
minutes, 60 minutes, 90 minutes, and 120 minutes after which the enzymatic reaction was
stopped by the addition of 2.5ml of TRIzol reagent.
RNA Extraction
RNA was extracted from digested homogenate with TRIzol reagent according to the
manufacturer’s instructions. The resulting RNA pellet was then resuspended in 20µl of
RNase-free water. The RNA concentration of each sample was measured using a
NanoDrop 2000 (Thermo Scientific). RNA was stored at -80°C for further analysis.
Complimentary DNA Library Preparation
First-strand cDNA synthesis was performed using 2µl of each RNA sample and
Superscript III reverse-transcriptase (Invitrogen) according to the manufacturer’s
instructions. The reaction was primed with tagged random septamers at 0.2µM (5’GCCGGAGCTCTGCAGATATCNNNNNNN-3’) and tagged oligo-dT primers at 0.2µM
(5’-GCCGGAGCTCTGCAGATATCTTTTTTT-3’) (Allander et al., 2005; Blomström et
69
al., 2010). Second-strand cDNA synthesis was performed by incubating the sample at
37°C for 1 hour with 1µl of the Klenow fragment (New England Biolabs).
PCR Amplification
GoTaq green master mix (Promega) was used to amplify cDNA with primers specific
to Actin and DWV, IAPB, SBV, and BQCV. Individual reactions were prepared with
1µl of cDNA and forward and reverse primers, each at a concentration of 0.4µM.
Reactions were subjected to one of three thermal (Table 2) depending on the annealing
temperature of the primer. Primer sequences and corresponding thermal protocol are
summarized in Table 1.
Visualization of PCR product
PCR products were loaded on a 1.5% agarose gel with a 100bp DNA ladder and
separated by electrophoresis.
2. Results and Discussion
This technique was being explored as a method to enrich an RNA preparation for
viral RNA while removing the excess mRNA and ribosomal RNA. It has been used
successfully for metagenomic studies using deep sequencing to identify novel viral
pathogens (Blomström et al., 2010). RNase A is an exonuclease specific to singlestranded RNA. Encapsidated viral RNA should be resistant to RNase A degradation
because the protein capsid can protect the RNA genome from RNase A degradation. In
contrast, free mRNA and ribosomal RNA is susceptible to degradation by RNase A.
A honey bee homogenate was prepared and passed through a 0.2µm cellulose filter to
remove bacteria and other contaminates. The homogenate was then divided into aliquots
and digested with DNase I (to remove genomic DNA) and RNase A (to remove mRNA
and rRNA) for varying amounts of time. Aliquots from the same initial homogenate
were used to ensure that each sample had the same viruses at the same concentrations
before digestion. The RNA was then extracted from each digested homogenate and the
concentration of each RNA sample was measured on a spectrophotometer (Table 12).
RNA was then tested for the presence of viral RNA using an RT-PCR assay (Figure 20).
70
Table 12: Concentration of RNA extracted from viral homogenate digested with RNase A
and DNase I for varying amounts of time. RNA was dissolved in a total of 20µl of RNase
free water.
Digestion Time (min) RNA Concentration (ng/µl)
0
1422.7
15
611
60
92
90
138.9
120
99.4
Figure 20: Susceptibility of actin mRNA and several honey bee Picornavirles viruses to
ribonuclease A and deoxyribonuclease I digestion.
As expected, the concentration of RNA extracted generally decreased with a longer
digestion time. It decreased between the 0 and 15 minute digestion and between the 15
and 60 minute digestion but remained similar for subsequent digestion times (Table 12).
71
This suggests that the nucleases are effectively digesting some but not all of the RNA and
some RNA is resistant to digestion. Not surprisingly, Actin mRNA was found in the
undigested homogenate but was not present in the digested homogenate. DWV, SBV,
and BQCV were all found in the undigested samples, however, only IAPV and DWV
were found in the digested samples suggesting that DWV and IAPV are resistant to
RNase A digestion while SBV and BQCV are not. It was also observed that the smallest
amount of PCR product for both DWV and IAPV was found in the 120 minute digestion,
suggesting that viral RNA was present in a lower amount in this sample compared to less
digested samples. This is likely because these viruses are not completely resistant to
RNase A digestion. IAPV was only found in digested samples suggesting that the
concentration of IAPV relative to other RNA in the undigested samples is much lower
than in digested samples and that the nuclease digestion enriched for this virus and
increased its relative concentration above a certain detectability threshold.
It is unknown why these viruses have different susceptibilities to RNase A digestion.
One hypothesis is simply that the amount of virus present in the homogenate was much
greater for DWV and IAPV so they required a longer digestion time to be completely
removed. Another hypothesis is that the capsids differ in permeability to RNase or
stability which could be altered by the specific buffer composition. A third hypothesis is
that DWV and IAPV contain a specific factor which makes them resistant to RNase A
digestion. For example, Yang and colleagues (2004) demonstrated that a specific
protease-sensitive factor protects newly synthesized Hepatitis C viral RNA from nuclease
degradation and it is possible that such a factor could exist for DWV or IAPV as well. A
fourth hypothesis is that DWV and IAPV are localized in a cellular vesicle or other
protected cellular structure. The vesicle, not the capsid, was shielding the genome from
RNase A. If this is the case, this result gives insight into how the replication cycles and
the location of replication within the host viruses differ among these viruses. A final
hypothesis is that the viral genome only exists in the homogenate as a double-stranded
replicative intermediate. Since RNase A is specific to single-stranded RNA, this doublestranded structure would be resistant to degradation.
72
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